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INTRODUCTION |
The mitochondrial respiratory chain (complexes I-V) is the major
site of ATP production in eukaryotes. Recently it was recognized that
this organelle not only generates ATP, but also plays an important role
in apoptosis (for reviews see Refs. 1-3).
It is now clear that upon apoptotic stimulation mitochondria can
release several proapoptotic regulators, including cytochrome c (4), Smac/Diablo (5, 6), endonuclease G (7), and apoptosis-inducing factor (8), to the cytosol. These proapoptotic regulators will then activate cellular apoptotic programs downstream (for reviews see Refs. 1-3). The release of proapoptotic regulators is
further regulated by the translocation of Bcl-2 family proteins (9,
10). Although this evidence places mitochondria in the center of the
apoptotic signaling pathway, the role of mitochondrial respiratory
chain activity in apoptosis is still elusive.
Inhibition of the mitochondrial respiratory chain by rotenone has been
widely used to study the role of the mitochondrial respiratory chain in
apoptosis (11-13). Some recent evidence showed that rotenone, a
mitochondrial respiratory chain complex I inhibitor, could induce cell
death in a variety of cells (11-16). The importance of mitochondrial
respiratory chain complex I inhibitor-induced apoptosis was further
recognized with the finding that tumor necrosis factor (TNF)-
could
inhibit the mitochondrial respiratory chain at the mitochondrial
complex I site (14). However, contradictory reports showed that
rotenone could inhibit apoptosis in other systems (17-19).
It has been suggested that
ROS1 play an important role
in apoptosis, and several groups have shown that molecules that
stimulate formation of ROS can result in apoptosis (20, 21) and a
process inhibited by antioxidants (22, 23). Others reported production of ROS by a wide range of apoptotic stimuli, including TNF, ceramide, staurosporine, and UV radiation (24-27). The mitochondrial respiratory chain is one of the most important sites of ROS production under physiological conditions (28-30), and it has been long suspected that
mitochondrial ROS play an important role in apoptosis. The mitochondrial-derived ROS are vital not only because mitochondrial respiratory chain components are present in almost all eukaryotic cells, but also because the ROS produced in mitochondria can readily influence mitochondrial function without having to cope with long diffusion times from the cytosol. Two sites in the respiratory chain,
complex I and complex III, have been suggested to be the major ROS
source (31-33). Based on stoichiometrical calculation, superoxide was
suggested as the primary product, with hydrogen peroxide as the
secondary product (34). Mitochondrial-derived ROS could be modified
when the mitochondrial respiratory chain was interrupted under
pathological conditions or by respiratory chain inhibitors (31, 33).
Early reports showed that the complex I inhibitor rotenone and the
complex b-c1 inhibitor antimycin could stimulate superoxide and
hydrogen peroxide formation on submitochondrial particles (31, 35).
However, results of rotenone-induced mitochondrial ROS production
measured at the cellular level appeared inconsistent with conflicting
results reporting that rotenone could elevate cellular ROS production
in some cases (11, 36) while inhibiting cellular ROS production in
others (17, 37, 38).
The present investigation studied the mechanism of rotenone-induced
apoptosis. Our data show that rotenone can induce mitochondrial ROS
production and that rotenone-induced mitochondrial ROS production is
closely related to rotenone-induced apoptosis.
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MATERIALS AND METHODS |
Cell Culture and Reagents--
Hoechst 33342, propidium iodide
(PI), and anti-cytochrome oxidase subunit VI antibody were obtained
from Molecular Probes, Eugene, OR. Hydroethidine was obtained from
Polysciences, Warrington, PA. Anti-human caspase 3 antibody and
anti-human cytochrome c antibody were obtained from Santa
Cruz Biotechnology, Inc., Santa Cruz, CA. All other reagents, unless
otherwise stated, were obtained from Sigma Chemical Co.
The human promyelocytic leukemia cell line HL-60 was obtained from ATCC
(Manassas, VA). Cells were cultured in RPMI 1640 medium supplemented
with 10% fetal calf serum and 2 mM
L-glutamine.
HL-60 cells lacking mitochondrial DNA (
°) were generated by
growing HL-60 cells in RPMI 1640 medium supplied with 10% fetal calf
serum, 2 mM L-glutamine, 1 mM
pyruvate, 50 µg/ml uridine, 25 mM glucose, and 50 ng/ml
ethidium bromide for 8 weeks as previously described (39). After
selection, the cells were grown in the same medium without ethidium
bromide. Oxygen consumption was measured with a Clark-type oxygen
electrode, and no oxygen uptake was observed for
° HL-60 cells.
The human fibrosarcoma cell line HT-1080 was maintained in Dulbecco's
modified Eagle's medium containing 10% heat-inactivated fetal calf
serum and supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate, and 100 units/ml penicillin.
Construction of a stable HT1080 cell line overexpressing magnesium
superoxide dismutase (Mn-SOD) was previously described in detail
(40). HT1080 cells stably transfected with Mn-SOD were maintained in medium that included 1 mg/ml G418 in addition to the above-mentioned supplements. All cell lines were cultured at 37 °C with 5%
CO2.
Respiration Measurement--
Oxygen consumption was measured
with a Clark oxygen electrode (model 5300: Yellow Spring Instrument
Co., Yellow Spring, OH) as described before (41). Briefly, 1 × 107 HL-60 cells were treated with various concentrations of
rotenone for 30 min. Cells were then collected and resuspended in a
medium containing 0.3 M mannitol, 10 mM
potassium HEPES (pH 7.4), 5 mM potassium phosphate (pH
7.4), and 1 mM MgCl2. The cells were injected into a respiration chamber that was then sealed. The total volume of
the respiration chamber was 1.6 ml. Respiration was then measured and
calculated as the rate of change in the oxygen concentration, assuming
the initial oxygen concentration to be 6.8 mg/liter. Cell respiration
was converted to percentage of control.
ATP Determination--
For ATP measurement, a commercially
available luciferin-luciferase assay kit was used. Briefly, HL-60 cells
were treated with various concentrations of rotenone for 24 h and
then collected in 1-ml Eppendorf tubes. After a single wash with
ice-cold PBS, cells were lysed with the somatic cell ATP-releasing
reagent provided by the kit. Luciferin substrate and luciferase enzyme
were added and bioluminescence was assessed on a Perkin Elmer 3B
spectroflurometer. Whole-cell ATP content was determined by running an
internal standard. The cellular ATP level was converted to percentage
of untreated cells (control).
Measurement of Hydrogen Peroxide Production by Intact
Mitochondria--
Intact mitochondria were isolated from cultured
HL-60 cells. HL-60 cells (1 × 109 cells) were
collected by centrifugation at 500 × g for 5 min. After one wash with ice-cold Ca2+, Mg2+-free
PBS at 250 × g, the cells were resuspended in 2 ml of
isolation buffer containing 250 mM sucrose, 10 mM HEPES (pH 7.4), and 1 mM EDTA. Cells were
homogenized in an ice-cold Dounce homogenizer with 20 strokes. Cell
disruption was confirmed by trypan blue staining. The disrupted cells
were centrifuged for 10 min at 600 × g at 4 °C.
Supernatant was collected and centrifuged at 15,000 × g, 4 °C, for 10 min, and the resulting pellet was
considered as crude mitochondria. The crude mitochondria were further
washed twice with the same isolation buffer as described above. The
resulting mitochondria pellet was resuspended in isolation buffer
without EDTA. Mitochondria suspensions were kept on ice, and all
experiments were performed within 5 h. Mitochondrial protein
concentration was determined by the biuret method (Bio-Rad) with bovine
serum albumin as the standard.
Mitochondrial hydrogen peroxide production was measured from the
increase of oxidized p-hydroxyphenylacetate (PHPA)
fluorescence by horseradish peroxidase (41, 43). Fluorescence of
oxidized PHPA (excitation 320 nm, emission 400 nm) was measured by a
Perkin Elmer 3B spectrofluorimeter. Mitochondria (0.5 mg of protein) were added to 4 ml of medium containing 0.3 M mannitol, 10 mM potassium HEPES (pH 7.4), 5 mM potassium
phosphate (pH 7.4), 1 mM MgCl2, 10 µg/ml
PHPA, 10 units of horseradish peroxidase, and the substrates for
various mitochondrial respiratory chain complexes. Mitochondrial
hydrogen peroxide production was determined by interpolation from the
standard curve generated by reagent hydrogen peroxide.
Measurement of Cellular Superoxide Production by Flow
Cytometry--
Measurement of cellular superoxide production was
performed as described previously, with some modifications (44). For
HL-60 cells, cells were treated with various concentrations of rotenone for 30 min. Cells were then collected and washed in Hank's balanced salt solution (HBSS) at 250 × g. Cells were
resuspended in HBSS containing 10 µM hydroethidine (HE)
and incubated at 37 °C for 10 min. For HT1080 fibrosarcoma cells,
cells were treated with rotenone for 30 min, the medium was then
removed, and cells were washed once with HBSS. Cells were then
incubated HBSS containing 0.25% trypsin for 5 min. Cells were then
resuspended in HBSS containing 10 µM hydroethidine (HE),
and incubated at 37 °C for 10 min. All cell suspensions were placed
into 12 × 75-mm tubes for assay. Flow cytometry studies were
carried out on a Beckman-Coulter XL flow cytometer. Ethidium
fluorescence was collected using a 610-nm long-pass filter.
DNA Fragmentation--
The DNA fragmentation assay was performed
according to the method described previously with some modifications
(45). HL-60 cells were treated with various concentrations of rotenone
in the presence or absence of antioxidants. Cells (2 × 107) were washed once with PBS (4 °C, pH 7.4) and
collected by centrifugation at 250 × g for 5 min. The
pellet was then treated with 0.5 ml of lysis buffer (10 mM
Tris-HCL, pH 7.4, 10 mM EDTA, 0.5% sodium dodecyl sulfate)
for 10 min on ice. After treatment with RNase A (final concentration,
100 µg/ml) for 1 h at 37 °C, the cells were incubated at
50 °C for 4 h in the presence of 100 µg/ml proteinase K. DNA
was precipitated by addition of 50 µl of 3 M sodium
acetate (pH 5.2) and 1 ml of cold (4 °C) 100% ethanol to the
solution. DNA was then collected and dissolved in TE buffer (10 mM Tris pH 8.0, EDTA 1 mM). For analysis,
10-20 µl of DNA was loaded on a 1.2% agarose gel containing 10 µg/ml ethidium bromide. Electrophoresis was performed in 0.5 × Tris borate-EDTA buffer (18 mM Tris-base (pH 8.0), 18 mM boric acid, and 1 mM EDTA) at 70 V for
2 h. DNA was visualized under ultraviolet light and photographed.
Subcellular Fractionation--
For HL-60 cells, 2 × 106 were collected by centrifugation at 250 × g for 5 min. For HT1080 cells, cells were collected by
trypsinization and then centrifuged (250 × g) for 5 min. All cells were then resuspended in buffer containing 250 mM sucrose, 10 mM HEPES, and 1 mM
EDTA, pH 7.4, 4 °C. Digitonin (400 ng/ml) was then added to the cell
suspension and lysis of cells confirmed by examining the entry of
trypan blue into the cytosol. The cell suspension was centrifuged at
5,000 × g for 10 s at 4 °C to pellet the
unbroken cells. The resulting supernatant was centrifuged at
12,000 × g for 10 min at 4 °C. The final
supernatant was considered as the cytosol fraction, and the final
pellet was used as the mitochondrial fraction.
Whole Cell Extraction--
For HL-60 cells, ~1 × 107 cells were harvested and washed once with ice-cold PBS,
resuspended in 1 ml of ice-cold lysis buffer containing 1% Nonidet
P-40, 20 mM Tris-HCl (pH 8.0), 10% glycerol, 137 mM NaCl, 2 mM EDTA, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, 1 mM dithiothreitol, and 1 mM sodium
orthovanadate, and incubated on ice for 30 min. After centrifugation at
12,000 × g for 10 min at 4 °C, cell lysates were
transferred to fresh tubes and stored at
80 °C. For HT1080 cells,
cells at 80-90% confluence in 100-mm culture dishes were washed once
with HBSS and placed on ice. In each culture dish, 400 µl of lysis
buffer was added. After 10 min of incubation, the solubilized proteins
were centrifuged at 12,000 × g in a microcentrifuge (4 °C) for 10 min, and the supernatants were stored at
80 °C.
Western Blotting--
Cell homogenates (50 µg of protein) were
fractionated by SDS-PAGE on a 15% acrylamide gel. Bands of proteins
were then transferred to a polyvinylidene difluoride (PVDF) membrane
(Bio-Rad). The PVDF membrane was blocked by PBS containing 5% milk
overnight at 4 °C and then incubated with antibodies against either
cytochrome c, caspase 3,
-actin, or cytochrome oxidase
subunit VI for 3 h at room temperature. After an additional 1-hour
incubation with horseradish peroxidase-conjugated secondary antibodies,
the binding of antibodies to the PVDF membrane was detected with an
enhanced chemiluminescence Western blotting analysis (Amersham Biosciences).
Measurement of Apoptosis by Flow Cytometry and Confocal
Microscopy--
Degradation of DNA was measured by flow cytometry and
confocal microscopy. For flow cytometry analysis, PI was used to detect DNA breakdown as described previously (46). Cells were collected and
fixed in suspension in 70% ethanol on ice, and then stored at
20 °C for at least 4 h. Cells were washed with 5 ml of HBSS, centrifuged again, and resuspended in 1 ml of HBSS. After addition of
0.2 ml of phosphate citrate buffer (0.2 M
Na2HPO4, 4 mM citric acid, pH 7.8),
cells were incubated at room temperature for 5 min before being washed
again and resuspended in HBSS containing 20 µg/ml PI and 10 µg/ml
RNase A. Cells were incubation in the dark at room temperature for 30 min. PI fluorescence was analyzed by flow cytometry.
Apoptosis was also measured by observing morphological changes in the
nuclear chromatin of cells detected by staining with 2 µg/ml Hoechst
33342 at room temperature for 15 min, followed by examination on a
Bio-Rad MRC 1024 confocal microscope.
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RESULTS |
Impairment of Mitochondrial Function in HL-60 Cells by
Rotenone--
Rotenone-induced mitochondrial dysfunction was
investigated by both respiration measurement and determination of
cellular ATP level. Fig. 1A
shows the dose-dependent response of rotenone-inhibited HL-60 cell respiration. Inhibition of cell respiration was detectable at rotenone levels as low as 10 nM. Below 100 nM, increasing concentrations of rotenone produced a rapid
decrease in the respiration of the cells. Above 100 nM, the
inhibition of respiration continued, but at a lower rate. Rotenone at
500 nM inhibited cell respiration by over 96%.

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Fig. 1.
Inhibitory effects of rotenone on cell
respiration and cellular ATP levels. A, HL-60 cells
were treated with various concentrations of rotenone for 30 min before
measurement of cell respiration. Respiration was calculated as
percentage of control. B, HL-60 cells were treated with
rotenone for 24 h and collected for whole cell ATP measurement as
described under "Materials and Methods." ATP values are expressed
as percent of control. All values are means ± S.E.,
n = 3.
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The curve of ATP inhibition by rotenone was similar to the respiration
curve (Fig. 1B). Increasing concentrations of rotenone below
100 nM result in a sharp decrease of cellular ATP level. Above 100 nM, cellular ATP level still decreased, but at a
much slower rate. Rotenone at 500 nM decreased cellular ATP
level to 64% of control. Higher concentrations of rotenone did not
further decrease the ATP level. By comparison, 10 µM
oligomycin inhibited cellular ATP level to 55% of control (data not shown).
Generation of Mitochondrial ROS by Rotenone in HL-60
Cells--
Production of mitochondrial reactive oxygen species by
HL-60 cells was estimated for both isolated mitochondria and cultured cells. For mitochondrial ROS measurement, a classical horseradish peroxidase-based PHPA oxidation method was used to detect hydrogen peroxide leaked from intact mitochondria when mitochondria were incubated with various respiratory chain substrates and inhibitors. As
reported by Hansford et al. (42), an inhibition of
mitochondrial hydrogen peroxide formation was observed in our system
when a higher concentration of PHPA was used. However, the
concentration at which we observed inhibition was much lower than that
reported by Hansford et al. (20 µg/ml versus 50 µg/ml). Below 20 µg/ml, inhibition of hydrogen peroxide formation
by PHPA was not observed (data not shown). Therefore, a final
concentration of 10 µg/ml PHPA was selected to be used in the current study.
Mitochondria from HL-60 cells were able to generate a low level of
hydrogen peroxide (Fig. 2A and
Table I) in the presence of mitochondrial
respiratory chain substrates, glutamate/malate for complex I and
succinate for complex II. The mitochondrial complex I inhibitor
rotenone increased glutamate/malate-supported hydrogen peroxide
formation. The concentration of rotenone capable of inducing
mitochondrial hydrogen peroxide formation was closely related to
the concentration that inhibited cell respiration. At low
concentrations, cell respiration levels were highly sensitive to
rotenone treatment. The respiration inhibition curve was very rapid with a nearly 80% decrease of cell respiration in the presence of 100 nM rotenone. Similarly, mitochondrial hydrogen
peroxide formation increased quickly within the same rotenone
concentration range. Respiration in cells treated with 0.5 µM rotenone was reduced to 4% of control levels. Higher
concentrations of rotenone produced greater inhibition but a much
slower decrease in cell respiration. Correspondingly, 0.5 µM rotenone stimulated the maximum rate of mitochondrial hydrogen peroxide production. Higher concentrations of
rotenone did not further elevate this rate.

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Fig. 2.
Effect of rotenone on ROS production.
A, mitochondria were isolated from HL-60 cells and incubated
with various concentrations of rotenone and glutamate/malate. Hydrogen
peroxide produced by mitochondria was measured as described under
"Materials and Methods." Mitochondrial hydrogen peroxide was
converted to nmol of hydrogen peroxide produced per min per mg of
mitochondrial protein. Values are means ± S.E., n = 3. B, intact HL-60 cells were treated with various
concentrations of rotenone for 30 min. Cells were then collected and
loaded with 10 µM hydroethidine for 10 min. ROS levels
were estimated by measuring ethidium fluorescence using flow cytometry.
C, cellular ROS production in 0 HL-60
cells treated with various concentrations of rotenone for 1 h
as indicated.
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Table I
Rotenone-induced mitochondrial hydrogen peroxide production
Mitochondria were isolated from HL-60 cells and incubated with various
respiratory chain substrates and inhibitors. Hydrogen peroxide produced
by mitochondria was measured as described under "Materials and
Methods." Mitochondrial hydrogen peroxide was converted to mnol of
hydrogen peroxide produced per min per mg of mitochondrial protein.
Values are means ± S.E., n = 3.
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Another mitochondrial respiratory chain inhibitor, antimycin, was also
able to stimulate mitochondrial hydrogen peroxide formation (Table I).
However, antimycin could elevate both glutamate/malate- and
succinate-supported mitochondrial hydrogen peroxide formation, while
rotenone was shown to increase only glutamate/malate-supported hydrogen peroxide formation.
To confirm that rotenone was able to induce mitochondrial ROS
production, conversion of the ROS-sensitive dye hydroethidine to
ethidium was used to measure cellular ROS level by flow cytometry (Fig.
2B). Using this technique, rotenone-induced ROS production was detected in single cells at the wholecell level. Rotenone was
shown to elevate cellular ROS levels in HL-60 cells. In comparison with
the dose-response curve of rotenone-induced mitochondrial hydrogen
peroxide formation, at the cellular level low concentrations of
rotenone (<100 nM) could merely elevate the cellular ROS
level. However, at high concentrations (
500 nM) rotenone
could significantly increase the cellular ROS level.
To further investigate whether rotenone-induced cellular superoxide
production was from mitochondria, HL-60 cells that lack mitochondrial
DNA (
0) were treated with various concentrations of
rotenone and then loaded with hydroethidine. Rotenone could not elevate
ethidium fluorescence in
0 HL-60 cells after treatment
for 1 h (Fig. 2C), although ethidium fluorescence
increased when normal HL-60 cells were treated with rotenone for the
same length of time (Fig. 2C).
Rotenone-induced Apoptotic Cell Death in HL-60 Cells--
Using
HL-60 cells as a model, we studied the effect of rotenone on cell
death. After treatment with 1 µM rotenone for 36 h, ~50% of the HL-60 cells showed chromatin condensation and nuclear fragmentation under confocal microscopy using Hoechst 33342 as DNA
stain (Fig. 3). Cell cycle analysis based
on flow cytometry was used to quantitatively estimate rotenone-induced
apoptotic cell death. Gating of the subdiploid cell population on a
linear scale excluded cells with extensive DNA degradation (typical of necrosis) to distinguish apoptotic cells from necrotic cells (45). Consistent with confocal microscope results, flow cytometry analysis revealed an increase in the subdiploid population after treatment with
1 µM rotenone for 36 h (55.3% compared with 6.7%
of control). Agarose gel electrophoresis studies showed that 0.5-1
µM rotenone treatment resulted in a typical
apoptotic-style DNA fragmentation with 30/50 kbp and 200/1000 bp
oligonucleosomal fragments (Fig. 4A). In addition, the number
of subdiploid cells increased in a concentration-dependent
manner (Table II). Rotenone (0.5 µM) for 36 h induced ~50% of cells to be
subdiploid. Higher concentrations of rotenone did not produce more
subdiploid cells than were induced by 0.5 µM rotenone
(Table II).

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Fig. 3.
Effect of rotenone on DNA breakdown in HL-60
cells. HL-60 cells were incubated with either Me2SO
(control) or 1 µM rotenone in
Me2SO for 36 h. Left panel, cells stained
with Hoechst 33342 and imaged on the confocal microscope.
Right panel, cells fixed and stained with PI as described
under "Materials and Methods." The number of apoptotic cells was
estimated by gating on the subdiploid population. Cells with extensive
DNA breakdown were not considered as apoptotic cells and were gated
out. Numbers on each gate show the percentage of apoptotic cells in
that histogram. Data are representative of three separate
flow cytometry analyses.
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Fig. 4.
Rotenone-induced DNA laddering and effect of
caspase inhibitors on rotenone-induced apoptosis. A,
HL-60 cells were treated with different concentrations of rotenone for
36 h. Genomic DNA was isolated and run on a 1% agarose gel and
visualized by ethidium bromide staining. Results are representative
from three separate experiments. B, cells were pretreated
with caspase inhibitors (Z-VAD and DEVD) for 30 min. Rotenone (1 µM) was then added to the medium. After
36 h, cells were collected, and the apoptotic cell number was
estimated by flow cytometry as described under "Materials and
Methods." Significant difference from control group at *,
p < 0.05. C, HL-60 cells or
0 HL-60 cells were incubated with either
Me2SO (control) or 1 µM rotenone
in Me2SO for 36 h. Cells were then collected, and the
apoptotic cell number was estimated by flow cytometry as described
under "Materials and Methods." Significant difference from control
group at *, p < 0.05.
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Table II
Rotenone-induced apoptosis in HL-60 cells
HL-60 cells were incubated with either Me2SO (control) or
various concentrations of rotenone in Me2SO for 36 h.
Cells were then fixed and stained with PI as described under
"Materials and Methods." The number of apoptotic cells was
estimated by gating on the subdiploid population. Cells with extensive
DNA breakdown were not considered as apoptotic cells and were gated
out. Data were represented as percentage of apoptotic cells in the
whole cell population. All values are means ± S.E.,
n = 3.
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The effects of two caspase inhibitors, the broad caspase inhibitor
z-VAD and the specific caspase 3 inhibitor DEVD-CHO (Bachem, King of
Prussia, PA), on rotenone-induced apoptosis were studied (Fig.
4B). Both reagents significantly inhibited rotenone-induced apoptosis, suggesting the involvement of caspases in rotenone-induced apoptosis. In
0 HL-60 cells, rotenone treatment could
cause only around 20% of cells to be apoptotic, much less than the
apoptotic cells in normal HL-60 cells (around 52%). We then examined
the biochemical properties of rotenone-induced apoptosis. Release of
cytochrome c from mitochondria to the cytosol and
subsequently activation of caspase 3 were considered as the biochemical
hallmarks of apoptosis. Western blotting results revealed that after
18 h of rotenone (0.5-1 µM) treatment,
mitochondrial cytochrome c decreased while cytosol
cytochrome c increased, indicating a typical release of
cytochrome c from mitochondria to the cytosol (Fig.
5A). The amount of cytochrome
c released to the cytosol increased in a
time-dependent manner. After a 30-hour treatment, cytochrome c release reached a plateau, and no further
increase of cytochrome c release was observed. Caspase 3 activation was also investigated (Fig. 5B). Caspase 3 was
activated at nearly the same time as cytochrome c release.
Similarly, after rotenone treatment for 18 h, cleavage of
pro-caspase 3 to release active caspase was observed. Maximum caspase 3 activity occurred after 30 h.

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Fig. 5.
Rotenone-induced cytochrome
c release and caspase 3 activation in HL-60
cells. HL-60 cells were treated with rotenone (0.5 or 1 µM) for various times as indicated. A,
cytosolic cytochrome c and mitochondrial cytochrome
c were determined by immunoblot analyses. B,
whole-cell lysate was obtained as described under "Materials and
Methods." Caspase 3 activation was determined by immunoblot analysis
using caspase 3 antibody to detect both pro-caspase 3 and active
caspase 3. -Actin level was used to confirm the equal loading of
samples.
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Inhibition of Rotenone-induced Apoptotic Cell Death by
Antioxidants--
To study the role of rotenone-induced production of
mitochondrial reactive oxygen species in apoptosis, the effects of
antioxidants on rotenone-induced apoptosis were examined. Three
classical antioxidants, glutathione, NAC, and vitamin C, were used in
the current investigation. All three antioxidants decreased the number
of subdiploid cells induced by rotenone treatment at concentrations
that are commonly used in other studies
(Figs. 6 and 7). Among these
antioxidants, vitamin C was shown to be the most potent.
Rotenone-induced apoptosis was almost completely inhibited by vitamin C
(Figs. 6E and 7A). DNA laddering produced by
rotenone treatment was also inhibited by the addition of antioxidants
(Fig. 7B), confirming the inhibition of rotenone-induced DNA
breakdown by these antioxidants.

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Fig. 6.
Inhibition of rotenone-induced DNA breakdown
by antioxidants. HL-60 cells were pretreated with either 15 mM glutathione, 15 mM
N-acetylcysteine, or 15 mM vitamin C for 30 min.
Either Me2SO (control) or 1 µM
rotenone in Me2SO was added to the medium. After 36 h,
cells were collected and apoptosis was detected by flow cytometry as
described under "Materials and Methods." Numbers on each gate show
the number of apoptotic cells in that histogram. Each
histogram is representative of three separate experiments.
From top to bottom: A, control (with
Me2SO); B, rotenone, 1 µM;
C, rotenone, 1 µM with 15 mM
glutathione; D, rotenone, 1 µM with 15 mM N-acetylcysteine; E, rotenone, 1 µM with 15 mM vitamin C.
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Fig. 7.
Inhibition of rotenone-induced DNA breakdown
and DNA laddering by antioxidants. HL-60 cells were treated with
either 15 mM glutathione, 15 mM
N-acetylcysteine, or 15 mM vitamin C for 30 min.
Either Me2SO (control) or 1 µM
rotenone in Me2SO was added to the medium. A,
after 36 h, cells were collected, and apoptosis was detected by
flow cytometry. Data represent percentage of apoptotic cells in the
whole cell population. Data are means ± S.E., n = 3. B, after 36 h, cells were also collected and genomic
DNA was isolated and run on a 1% agarose gel and visualized by
ethidium bromide staining. Results are representative from three
separate experiments.
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Both cytochrome c release and caspase 3 activation were
found to be inhibited by all three antioxidants (Fig.
8). Similarly, vitamin C was shown to be
the most potent among the three. Treatment of cells with 15 mM vitamin C blocked cytochrome c release and caspase 3 activation almost entirely (Fig. 8).

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Fig. 8.
Inhibition of rotenone-induced cytochrome
c release and caspase 3 activation by
antioxidants. HL-60 cells were treated with either glutathione,
N-acetylcysteine, or vitamin C for 30 min. Either
Me2SO (control) or 1 µM rotenone
in Me2SO was added to the medium. After 36 h, cells
were collected. A, cytosolic cytochrome c and
mitochondrial cytochrome c were determined by immunoblot
analyses. B, whole-cell lysate was obtained as described
under "Materials and Methods." Caspase 3 activation was determined
by immunoblot analysis using caspase 3 antibody to detect both
pro-caspase 3 and active caspase 3. -Actin level was used to confirm
the equal loading of samples.
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Inhibition of Rotenone-induced Cell Death by Mn-SOD
Overexpression--
To confirm that mitochondrial reactive oxygen
species play an important role in rotenone-induced apoptosis, an
Mn-SOD-overexpression cell model was constructed using HT1080
fibrosarcoma cell lines. Two cell lines, CMV (representing HT1080 cells
transfected with empty vector) and HT15 (representing HT1080 cells
transfected with vector containing Mn-SOD, 15-fold increase in Mn-SOD
levels after transfection), were used in the current study. Cellular ROS levels were examined by flow cytometry using hydroethidine as the
ROS indicator (Fig. 9A). Basal
ethidium fluorescence of HT15 cells was lower than that of CMV cells.
Both CMV and HT15 cells showed increase in ethidium fluorescence after
1 µM rotenone treatment for 30 min, showing that rotenone
elevated cellular ROS level in both cell lines. However, the
increase of ethidium fluorescence in HT15 cells was about 50% lower
than that in CMV cells.

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Fig. 9.
Effect of Mn-SOD overexpression on
rotenone-induced ROS production, cytochrome c release,
and caspase activation. A, control HT1080 cells
(CMV) and HL-60 cells overexpressing Mn-SOD (HT15 = 15-fold elevation of Mn-SOD level) were treated with 1 µM
rotenone for 30 min. Cells were then loaded with 10 µM
hydroethidine. Cellular ROS levels were estimated by measuring ethidium
fluorescence using flow cytometry. B, CMV and HT15 cells
were treated with either Me2SO or 1 µM
rotenone in Me2SO for 36 h. Cytosolic cytochrome
c and mitochondrial cytochrome c were determined
by immunoblot analyses. C, caspase 3 activation was
determined by immunoblot analysis by caspase 3 antibody to detect both
pro-caspase 3 and active caspase 3. -Actin level was used to confirm
the equal loading of samples.
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The effects of rotenone on cytochrome release and caspase 3 activation
were also studied in both cell lines. Fig. 9B shows the
release of cytochrome c after rotenone treatment. CMV cells and HT15 cells showed similar basal cytochrome c expression
in mitochondria. Upon treatment with 1 µM rotenone for
24 h, a great portion of the mitochondrial cytochrome c
of CMV cells was released from mitochondria to the cytosol. However,
mitochondrial cytochrome c was largely maintained in
mitochondria of HT15 cells. Overexpression of Mn-SOD also inhibited
rotenone-induced caspase 3 activation on HT1080 cells. Fig.
9C indicates that after rotenone treatment for 24 h,
pro-caspase 3 was largely cleaved in CMV cells. However, in HT15 cells
the cleavage of pro-caspase 3 was much less.
Cell cycle analysis by flow cytometry was used to quantitatively
estimate the number of apoptotic cells after rotenone treatment in both
cell lines. Both untreated CMV and HT1080 cells showed a typical
G1S/G2M distribution on flow cytometry (Fig.
10, A and B).
After treatment with rotenone (1 µM) for 24 h, more
than 50% of CMV cells moved to the subdiploid area, which is typical
for apoptotic cells. However, only 20% of the HT15 cells were in the subdiploid area (Fig. 10, C and D).

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Fig. 10.
Effect of Mn-SOD overexpression on
rotenone-induced apoptosis. CMV and HT15 cells were treated
with either Me2SO or 1 µM rotenone in
Me2SO for 36 h and then collected. Apoptosis was
detected by PI staining using flow cytometry. Numbers on each gate
represent the number of apoptotic cells in that histogram. Each
histogram is representative of three separate experiments.
From top to bottom: A, CMV control
(with Me2SO); B, HT 15 control (with
Me2SO); C, CMV with 1 µM rotenone
treatment; D, HT 15 with 1 µM rotenone
treatment.
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DISCUSSION |
In this study we demonstrated that the ability of the
mitochondrial respiratory chain complex I inhibitor rotenone to induce programmed cell death is closely related to its ability to induce mitochondrial ROS production. Previously, rotenone had been reported to
enhance glutamate/malate-supported mitochondrial reactive oxygen species production on both bovine heart submitochondrial particles and
rat heart intact mitochondria (31, 42). Our result confirmed that in
isolated HL-60 cell mitochondria, rotenone induced mitochondrial ROS
production through a similar mechanism. The induction of mitochondrial ROS production by rotenone had frequently been attributed to the ability of rotenone to block mitochondrial respiratory chain complex I,
thereby increasing the formation of ubisemiquinone, the primary electron donor in mitochondrial superoxide generation. This also appeared to be the case for HL-60 mitochondria because the
concentrations of rotenone that induced mitochondrial ROS corresponded
to the concentrations of rotenone that inhibited cell respiration. In addition, rotenone at 0.5-1 µM resulted in a maximum
decrease of cellular ATP level (~63% of control). The respiratory
uncoupler oligomycin, which shuts down electron input from both complex I and complex II, resulted in only 8% more inhibition on cellular ATP
level (~55% of control), indicating mitochondrial complex I
substrates are the major substrates for the mitochondrial respiratory chain in HL-60 cells. For HL-60 cells, these results suggest that once
rotenone inhibited mitochondrial respiratory chain complex I, the
accumulation of pyruvate/malate in the mitochondria could create a
condition that may well be similar to the isolated mitochondrial model.
The whole-cell-level ROS measurement by flow cytometry using
hydroethidine confirmed our findings on isolated mitochondria. Rotenone
induced cellular ROS production dose-dependently. Rotenone could not induce cellular ROS production on
0 HL-60
cells, further confirming that rotenone-induced cellular ROS originated
from mitochondria. At the cellular level, rotenone could enhance
cellular ROS level in a similar pattern through a slightly different
mechanism. Similar to the results from isolated mitochondria,
increasing concentrations of rotenone greater than 0.5 µM
produced further but much slower increases in cellular ROS production.
However, at lower concentration (100 nM), rotenone was able
to increase cellular ROS level only slightly, whereas in the isolated
mitochondria model, this concentration could induce a mitochondrial ROS
production with close to 50% of maximum ROS production (for rotenone
at 1 µM). The possible explanation is that at the
cellular level, the cellular antioxidant system may have a greater
impact on the decrease of ROS production.
Rotenone has been reported to cause cell death in a variety of cell
lines (11-16). However, whether this cytotoxicity leads to apoptosis
or necrosis may depend upon cell type. Our results showed that in HL-60
cells rotenone induced cell death through an apoptotic mechanism. Cell
cycle analysis showed a typical apoptotic subdiploid population after
rotenone treatment. Chromatin condensation and DNA breakdown were also
clearly observable by confocal microscopy. DNA laddering presented as a
typical apoptotic DNA cleavage into 50 kbp and 200/1000 bp
oligonucleosomal fragments. In addition, Western blots identified
release of cytochrome c from the mitochondrial compartment
to the cytosol after 0.5-1 µM rotenone treatment. Caspase 3 activation was also demonstrated. All these data indicated that in HL-60 cells rotenone-induced cell death occurs mainly via an
apoptotic mechanism as opposed to necrosis. A possible mechanism might
be that rotenone inhibits the mitochondrial respiratory chain at the
complex I site, decreasing the cellular ATP level; however, the
dependence of pyruvate/malate-supported ATP production varies in
different cell types. It has already been well established that ATP can
act as a switch between apoptosis and necrosis (47-49). A depleted
cellular ATP level (usually to around 30% of control) has been shown
to inhibit apoptosis (47-49). Therefore, in cell lines highly
dependent on pyruvate/malate-supported ATP production, rotenone
treatment might drastically decrease the cellular ATP level, which
could switch cell death from apoptotic to necrotic. However, this is
not the case in HL-60 cells, in which rotenone decreased cellular ATP
to a moderate level (63%). Both our results and other reports show
that even oligomycin decreases the cellular ATP level in HL-60 cells to
around 50-60% of control after 24 h treatment (50). In several
other cell lines, such as Jurkat, HeLa, or thymocyte, the same
concentrations of oligomycin can decrease the cellular ATP level to
less than 20% of control level (47-49). While unlikely to be
responsible for these results, the possible impact of high
concentrations of rotenone on cellular glycolytic pathways has not been
entirely excluded.
Our results strongly suggest that in HL-60 cells, induction of
mitochondrial ROS could well be the most significant mechanism of
rotenone-induced apoptosis. The concentrations of rotenone that induced
apoptosis were in the same range as the concentrations that induced
mitochondrial ROS production. Rotenone-induced apoptotic cells in
0 HL-60 cells were much fewer than those in normal HL-60
cells. Several antioxidants (glutathione, vitamin C, and
N-acetylcysteine) were shown to inhibit rotenone-induced
cytochrome c release, caspase 3 activation, and DNA
breakdown in HL-60 cells. Induction of mitochondrial ROS by rotenone
also occurred very early (<1 h) compared with cytochrome c
release and caspase 3 activation, suggesting that ROS production is
upstream of these apoptotic events.
Results from Mn-SOD-overexpressing HT1080 cells confirmed our
conclusion that rotenone induced apoptosis via an induction of
mitochondrial ROS production. Mn-SOD is the superoxide dismutase that
localizes in mitochondria. Mn-SOD is a primary component of the
cellular defense system against oxidative toxicity since superoxide can
react with hydrogen peroxide to generate singlet oxygen and hydroxyl
radicals, which are more toxic than superoxide and hydrogen peroxide
(51). In mitochondria, manganese superoxide dismutase is the major
enzyme responsible for converting superoxide to hydrogen peroxide
(52-54). Overexpression of Mn-SOD inhibited rotenone-induced increase
of cellular ROS, confirming that rotenone-induced cellular ROS
production in HT1080 cells was from mitochondria. Consistent with our
findings in HL-60 cells, overexpression of Mn-SOD also inhibited
rotenone-induced cytochrome c release, caspase 3 activation,
and DNA breakdown.
The mechanism of rotenone-induced apoptosis is still elusive, and
obscured by the occurrence of several concomitant events, including
shutdown of the electron transfer through respiratory chain complex I,
decreasing cellular ATP level, increasing mitochondrial ROS production,
and decreasing mitochondrial membrane potential. Previously, the
decrease of mitochondrial membrane potential and the opening of the
mitochondrial permeability transition pore, but not ATP reduction, have
been shown to be involved in rotenone-induced apoptosis (11-13).
However, the role of rotenone-induced mitochondrial ROS production has
not been fully investigated. Results from the current study identified
mitochondrial ROS as playing a key role in rotenone-induced apoptosis.