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Department of Molecular and Cellular Physiology, Louisiana State University Medical Center, Shreveport, Louisiana 71130-3932, USA
1Correspondence: Department of Molecular and Cellular Physiology, Louisiana State University Medical Center, 1501 Kings Hwy., Shreveport, LA 71130-3932, USA. E-mail: taw{at}lsumc.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: GSH/GSSG ratio • intestinal cell death • redox shifts • peroxidized lipids • intestinal oxidative stress • diamide and GSH oxidation
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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.
Apoptosis is a common mechanism that regulates cell death in
physiological processes like embryogenesis and normal tissue and organ
involution. Apoptosis can also be induced in cells by imposition of
external stresses such as bacterial toxins, heat shock, radiation, and
oxidative stress (2
3
4
5)
. Failure of apoptosis is
considered to contribute to the development of human malignancies,
including gut pathology such as cancer (6
, 7)
.
Peroxidized lipids is a class of dietary oxidants that can initiate
intestinal degenerative processes via generation of oxygen radicals
(8
, 9)
. Although previous studies have documented
cytotoxicity in association with consumption of lipid hydroperoxide
in vivo (10
11
12)
, the potential impact of
subtoxic levels of lipid hydroperoxides on intestinal integrity is
poorly understood. Bull et al. found that intrarectal
instillation of hydroperoxy and hydroxy fatty acids provoked
proliferative responses in colonic mucosa in rats (13)
.
Hara et al. found that rats given oxidized ethyl linoleate developed
mucosal hypertrophy of the large intestine (14)
. These
studies demonstrate that normal intestinal cell turnover can be
disrupted by lipid hydroperoxides and underscore the tumorigenic
potential of oxidized lipids. In a recent study (Gotoh, Y., Wang,
T. G., Rhoads, C. A., Fujimoto, K., and Aw, T. Y.,
unpublished results), we have demonstrated that subtoxic low levels of
lipid hydroperoxide (1–5 µM) induce phase transition of intestinal
cells from a quiescent to that of a proliferative state that is
mediated by peroxide-induced disruption of cellular redox balance. The
kinetics of oxidative shift in intestinal cell redox status in response
to hydroperoxide exposure was temporally linked to the expression of
intestinal cell proliferative markers and the induction of cell cycle
progression. In other studies, we found that lipid hydroperoxide
concentrations that are 100-fold higher than those that elicited
proliferative responses (0.1–0.2 mM) significantly injure CaCo-2 cells
(15)
. Although the imposition of a severe oxidant stress
often results in a cytotoxic biological end point, necrotic cell death
is not necessarily an obligatory end point of all oxidative stress
(16)
. The issue that oxidant-mediated induction of
oxidative shift in the cellular redox status can enhance proliferative
or apoptotic rather than necrotic responses has been suggested
(17
18
19)
. Indeed, shifting cellular control checkpoints in
the direction of reductant or oxidants during oxidative stress could
result in a cell that favors quiescence, proliferation, or apoptosis
(16
, 17)
.
The human colon carcinoma cell line CaCo-2 spontaneously exhibits
structural and functional characteristics of mature small bowel
enterocytes under standard culture conditions (20)
. In the
current study we used this cell model to test the hypothesis that
disruption of redox balance by subtoxic levels of lipid peroxide (5–25
µM) at levels higher than those that promote cell proliferation would
favor an apoptotic response. The specific objectives of this study were
to 1) determine the range of subtoxic lipid hydroperoxide
levels that induce CaCo-2 cell apoptosis, 2) define the
kinetics of oxidative shifts in redox status in response to
hydroperoxide challenge, and 3) determine the temporal
relationship between peroxide-induced redox imbalance and apoptosis.
Because the intestinal epithelium is constantly challenged by oxidants
like lipid hydroperoxides of either dietary or endogenous origins, the
study will provide important information on oxidant-mediated loss of
thiol/disulfide homeostasis and its role in regulation of intestinal
cell death.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell culture
Human carcinoma cell line CaCo-2 cells were obtained from the
American Type Culture Collection (Rockville, Md.) and seeded in 25
cm3 culture flasks. The cells were grown in DMEM
supplemented with 10% fetal calf serum (FCS), 1% nonessential amino
acids, 100 U/ml penicillin, and 10 mg/ml gentamicin. The cell cultures
were maintained in humidified atmosphere with 5%
CO2 95% air at 37°C. The culture medium was
changed every 3 days.
Preparation of lipid hydroperoxide emulsion
Lipid hydroperoxides (LOOH) were generated by air oxidation of
Menhaden oil for 5 days. Total LOOH content was quantified
spectrophotometrically by the thiobarbituric acid assay
(21)
and the oxidation products were identified by
high-performance liquid chromatography (HPLC) (15)
. LOOH
emulsions were prepared by sonicating 80 µmol oxidized oil and 570
µmol sodium taurocholate in 30 ml phosphate-buffered saline (PBS), pH
7.4, as described (22)
.
Cell incubations
Incubations were performed in 100 mm dishes at a cell density of
2 x 106. In dose-dependent studies, cells
were exposed to LOOH from 0–25 µM for 24 h, whereas in time
course studies cells were treated with 10 µM for various times from 0
to 48 h. In experiments to delineate the specific role for GSH or
GSSG, cells were treated with buthionine sulfoximine (BSO, 2 mM),
diamide (0.5 mM), or a combination of both agents. BSO is a potent
inhibitor of 
-glutamyl cysteine synthetase, the rate-limiting step
in GSH synthesis (23)
. Inhibition of GSH synthesis causes
a marked decrease in the total GSH pool but has a minimal effect on the
GSH/GSSG ratio. Diamide is a cell-permeant thiol oxidant that
specifically targets the thiols of GSH and free SH groups of proteins
(24)
. The action of diamide induces formation of disulfide
bonds via thiol-diamide intermediates (25)
, thereby
promoting the formation of GSSG or protein disulfide cross-link.
Consequently, the redox potential of the cell is shifted in favor of a
more oxidized state, which typically is reflected in an increase in the
GSSG to GSH ratio.
TUNEL (Tdt-mediated dUMP nick-end labeling) assay
The TUNEL assay measures fragmented DNA in apoptotic cells by
catalytically incorporating fluorescein-12-dUMP at the 3'-OH ends. The
TUNEL assay was carried out as described (Apoptosis Detection System
Kit, Promega). Briefly, cells (2x106) were
washed with PBS, fixed with 1% formaldehyde on ice, and permeabilized
with 70% ice-cold ethanol at -20°C overnight. The permeablized
cells were washed three times with PBS and DNA strand breaks were
labeled with fluorescein-12-dUMP by incubating cell pellets with
terminal deoxynucleotidyl transferase and nucleotide mix at 37°C. The
reaction was terminated by addition of 20 mM EDTA, and the cells were
incubated with propidium iodide (PI, freshly diluted to 5 µg/ml in
PBS) containing 250 µg of DNase-free RNase A. Cells with
fluorescein-12-dUTP-labeled DNA were quantified by flow cytometry.
Flow cytometric analysis was performed using a FACS Vantage flow cytometer (Becton Dickinson, San Jose, Calif.). The spectral overlap between the fluorochromes was determined by single-stained samples and was compensated for electronically. A minimum of 10,000 cells per sample were analyzed. Data analysis was carried out using Cell Quest software (Becton Dickinson) and was gated on pulse-processed PI signals to exclude doublets and large aggregates, using a multiparameter gate strategy. Cell debris was excluded from analyses based on significantly diminished light scatter (FSC, SSC) properties.
Quantification of DNA fragmentation and agarose gel
electrophoresis
CaCo-2 cells were grown in flasks at a density of 2 x
105 cells/flask in DMEM media supplemented with
10% FCS. After LOOH exposure, cells were lysed with 0.1% Triton X-100
and DNA fragmentation was determined by the diphenylamine method
(26)
. Results are expressed as the ratio of the amount of
nonfragmented (pelleted at 27,000 g) and fragmented DNA
(supernatant). Semiquantitative assessment of DNA fragments was
performed by agarose gel electrophoresis using 1.5% gel strength
containing 1 µg/ml ethidium bromide. Electrophoresis was performed
for 2 h at 70 V and DNA was visualized by UV fluorescence.
Determination of 8-hydroxy 2-deoxyguanosine (8-oxoG)
Oxidative damage to DNA was quantified by the formation of
8-oxoG (27)
. Total DNA was extracted from 3 x 10
6 cells with a chaotropic iodine protocol (WAKO
Chemicals, Kyoto, Japan) that minimizes ex vivo DNA
oxidation. Briefly, cells were homogenized on ice in 0.1M NaCl, 30 mM
Tris, pH 8.0, 10 mM EDTA, 10 mM ß-mercaptoethanol, 0.5% Triton-X-100
with six passes of a Teflon glass homogenizer. The nuclei fraction was
collected and incubated with RNase (50 U/ml RNase A and 100 U/ml RNase
T1), proteinase K (5 mg/ml) in 10 mM Tris-HCl, pH 8.0, 5 mM EDTA, and
1% sodium dodecyl sulfate (SDS) at 50°C to remove contaminating RNA
and protein. DNA was extracted from the 10,000 g supernatant
using the iodine protocol consisting of NaI treatment and extraction
with 40% 2-propanol and isopropyl alcohol. The extracted DNA was
incubated with 0.1 mg/ml nuclease P1 at 65°C, followed by 1 U/µl
alkaline phosphatase. The deoxyribonucleosides were purified (10,000
g) on an UltraFree Millipore Eppendorf Filtration system
with a 30,000 Da cutoff and separated on a 15 cm x 4.6 mm 3 mm
LC-18 DB Supelco column (Bellefonte, Pa.) using a 6% linear gradient
of methanol in 50 mM KH2PO4
buffer (pH 5.5) and a flow rate of 1 ml/min; 8-oxoG was detected using
an ESA electrochemical detector (0.6 volts); results are expressed as
the number of 8-oxoG residues per µg DNA.
Preparation of cell lysate for Western blot analysis of caspase 3
activity
Caspase 3 activation was determined by cleavage of the
procaspase CPP32 and its activity by the cleavage of its endogenous
substrate, PARP, using Western analyses as described previously
(28)
. At designated times, control and LOOH-treated CaCo-2
cell monolayers were washed twice with ice-cold PBS and harvested into
25 mM HEPES buffer, pH 7.5, containing 5 mM EDTA, 1 mM EGTA, 5 mM
MgCl2, 2 mM DTT, 10 µg each of pepstatin and
leupeptin, and 1 mM PMSF. The cell suspension was sonicated on ice and
the cell lysate was centrifuged at 160,000 g. The pellet was
solubilized (in 25 mM HEPES, pH 7.5 5 mM EDTA, 2 mM DTT, 1% Triton
X-100, 10 µg each of pepstatin, and leupeptin and 1 mM PMSF),
sonicated, and used for Western blotting with the PARP antibody.
Protein samples (50 µg) were resolved on 8% SDS-polyacrylamide gels
and transferred to nitrocellulose membranes. The membranes were blocked
overnight in 5% non-fat milk in PBS and 0.1% Tween-20 at 4°C, and
thereafter incubated with anti-CPP32 (1:1000 dilution) monoclonal
antibody at room temperature. The membranes were washed (PBS with 0.1%
Tween-20) and incubated with horseradish peroxidase-conjugated goat
anti-mouse immunoglobulin G (1:1500 dilution). The immune complexes
were visualized by ECL according to the manufacture’s protocol. The
membranes were then stripped and probed with anti-PARP (1:5000
dilution), then stripped a second time and probed with anti-actin
antibody (1:2000 dilution) to verify equal protein loading in each
lane.
Biochemical assays
Cell GSH and GSSG were determined by the HPLC method of Reed et
al. (29)
. Cells were treated with ice-cold 5% TCA,
followed by centrifugation to remove TCA-insoluble proteins. The acid
supernatant was derivatized with 6 mM iodoacetic acid and 1%
2,4-dinitrofluorobenzene to yield the S-carboxymethyl and
2,4-dinitrophenyl derivatives of GSH and GSSG. Separation of GSH and
GSSG derivatives was achieved on a 15 cm x 4.6 mm 10 µm
C18-reversed phase ion-exchange column. Cell toxicity was determined by
lactate dehydrogenase (LDH) leakage, an indicator of cytoplasmic
membrane damage. LDH activity (30)
was assayed in the
culture media supernatants (S) at designated times after LOOH
treatment. Total cellular LDH was determined in the lysate after
treatment of corresponding cell monolayers with 5% Triton X-100. The
percentage of LDH release was calculated from the ratio of enzyme
activities in the supernatants and cell pellets according to the
equation [S/(S+C)] x 100.
Statistical analysis
Data are expressed as mean ± SD from four to
five individual cell preparation. Unless otherwise indicated,
individual data points in the figures that display no error bars
represent SD smaller than the size of the symbol. Data were
evaluated by analysis of variance in which multiple comparisons were
performed by the method of least significant difference. P
< 0.05 was considered as significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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shows the effect of LOOH on CaCo-2 apoptosis as measured by the TUNEL
method (Fig. 1A, B
) or agarose gel electrophoresis (Fig. 1C
). The results show that the percent of TUNEL-positive
cells in untreated controls was 3.5 ± 1.2%; this percentage
increased to 24.6 ± 1.7% at 24 h after exposure of cells to
10 µM LOOH, consistent with oxidant-induced apoptotic cell death.
Resolving agarose gel electrophoresis of total cell DNA shows that LOOH
caused a dose-dependent increase in DNA fragmentation with significant
increases at 10 µM and 20 µM LOOH (Fig. 1C
, lanes 3 and
4). Moreover, the DNA fragments were discernible as bands that are
consistent with endonuclease-mediated cleavage of nuclear chromatin
into discrete oligonucleosome-length fragments of 180 bp characteristic
of apoptotic cells (31)
. We found that measurements of
apoptosis by percent DNA fragmentation were quantitatively similar to
those assayed by the TUNEL method.
|
To determine whether the concentrations of LOOH that promote apoptosis
cause cytotoxicity, we measured the release of cytosolic LDH as an
index of cell membrane damage. The results in Fig. 2
show that exposure of cells to LOOH concentrations between 1 µM and
25 µM resulted in a dose-dependent increase in apoptosis (10–30%).
In comparison, LDH release was low—between 0–10 µM LOOH—and was
significantly detected only at 25 µM LOOH (14%). These results
demonstrate that at low oxidant levels, there was a clear dissociation
of apoptosis from necrosis that was correlated with increased severity
of oxidant challenge. In subsequent experiments, we used 10 µM LOOH
for our studies to induce CaCo-2 cell apoptosis without the
complication of causing cell necrosis.
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Lipid hydroperoxide disrupts cellular redox balance, which is
temporally correlated with enhancement of apoptosis
The dose-dependent relationship between changes in cellular GSH
and GSH/GSSG with apoptosis is illustrated in Fig. 3
. Exposure of cells to different LOOH concentrations (0–25 µM) for
24 h caused marked changes in cell GSH. LOOH concentrations
between 10 µM and 25 µM significantly decreased GSH to 20% control
levels (Fig. 3A
). LOOH at 0.4 µM caused a small increase
in cellular GSH (Fig. 3A
), suggesting that mild oxidant
stress promotes a compensatory increase in GSH, consistent with our
previous observations (Gotoh, Y. et al., unpublished results). In
comparison, the GSH/GSSG ratio fell at all LOOH concentrations (Fig. 3B
). Maximal GSH loss and decreased GSH/GSSG ratio were
achieved at 10 µM and 25 µM LOOH and was correlated with maximal
cell apoptosis (see Fig. 2
), indicating that a disruption of redox
homeostasis occurs with apoptosis. Figure 4
illustrates the kinetic relationship between LOOH-induced changes in
cell GSH and GSH/GSSG ratio and apoptosis. The results show that
treatment of cells with 10 µM LOOH caused a marked and rapid
oxidation of GSH by 15–30 min. By 1 h, cell GSH level was 30% of
control and remained low throughout the 48 h experimental period.
An increase in GSSG paralleled the loss of GSH, causing a decrease in
the GSH/GSSG ratio that tracked with the kinetics of GSH decrease. In
comparison, the extent of apoptosis was low within the first 12 h
after LOOH exposure and thereafter significantly increased from 16 h (15%) to 48 h (30%) postoxidant treatment (Fig. 4)
. These
results further show that LOOH-induced loss of redox balance preceded
detectable DNA fragmentation by 12–14 h.
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LOOH causes activation of caspase 3
To verify that the early loss of GSH/GSSG balance is an early
event associated with initiation of the apoptotic cascade, we examined
the activation of caspase 3 by determining the cleavage of the
procaspase, CPP32. Under unstimulated conditions (i.e., without LOOH),
CPP32 was undetectable in CaCo-2 cell extracts (Fig. 5A
, lane 1). Notably, cells treated with 10 µM LOOH
consistently exhibited an initial significant increase in the
expression of CPP32 within 30 min (lane 2), which over time decreased
(lane 4) to control levels by 2 h (lanes 5), consistent with an
early expression of the proenzyme, followed by its cleavage to the
active caspase during peroxide exposure. To further examine
LOOH-induced activation of caspase, we determined the kinetics of
cleavage of its endogenous substrate, PARP. The results in Fig. 5B
show that control cells expressed one significant protein
band that corresponded to the native PARP enzyme of 116 kDa and a faint
85 kDa band that corresponded to the cleavage product (lane 1).
Exposure of cells to 10 µM LOOH resulted in an initial increase in
the level of the 85 kDa product at 15 min, which was probably due to
protein overloading (lane 2). The cleavage of native PARP to its 85 kDa
and 29 kDa products increased over time (lanes 3 and 5) and peaked at
2 h (lane 5). Taken together, the results with CPP32 expression
and PARP cleavage are consistent with activation of caspase 3 within
2 h after the induction of cellular redox shift after LOOH
challenge.
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Diamide-induced oxidation of GSH and decreased GSH/GSSG ratio link
redox imbalance to induction of apoptosis
To directly test the role of redox change in the initiation of
apoptosis and delineate the contribution of GSH/GSSG ratio from that of
GSH per se, we used two chemical treatment strategies to
induce specific changes in either GSH alone (using BSO) or in both GSH
and GSSG (using diamide). Treatment of cells for 24 h with BSO at
2 mM resulted in > 95% depletion of cell GSH, but with minimal
change in the GSH/GSSG ratio (Fig. 6B, C
). Under these conditions, the percent of apoptosis was
essentially unchanged from control values (5%, Fig. 6A
),
suggesting that induction of a severely depleted GSH pool alone without
changes in the GSH/GSSG redox status was without effects on apoptosis.
Notably, diamide treatment for 24 h did not alter the cellular GSH
or GSH/GSSG ratio (Fig. 6B, C
), yet apoptosis of CaCo-2
cells was significantly increased (18%, Fig. 6A
). The
combined treatment of BSO and diamide yielded significantly decreased
GSH and an unchanged GSH/GSSG ratio (Fig. 6C
), consistent
with inhibition of GSH synthesis by BSO without effects on the redox
ratio. Cell apoptosis was increased and was quantitatively similar to
that elicited by diamide alone (18%, Fig. 6A
). Exposure of
CaCo-2 cells to BSO or diamide or BSO plus diamide at the designated
concentrations were without cytotoxic effects as measured by LDH
leakage (data not shown).
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The findings with diamide suggest that either the cellular GSH/GSSG
status do not play a role in apoptosis, which would be inconsistent
with the studies using LOOH, or that early GSH/GSSG change was the
primary inciting event of the apoptotic cascade; once initiated, the
recovery of redox balance did not prevent the progression of apoptosis
to its biological end point at 24 h later. To test the latter
possibility, cells were sampled for determination of GSH and GSH/GSSG
status at 30 min or 24 h after diamide treatment. The results in
Fig. 7
show that exposure of cells to the thiol oxidant resulted in >
90% oxidation of GSH (Fig. 7A
) with a concomitant increase
in GSSG at 30 min (Fig. 7B
). By 24 h, both cell GSH and
GSSG recovered to pretreatment baseline values. These results are
consistent with a distinct dissociation of the kinetics of GSH/GSSG
change (within 30 min) from the apoptotic end point (24 h).
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To determine the threshold window of redox imbalance in initiating the
apoptotic process, we compared the time course of diamide-induced loss
and recovery of GSH and GSH/GSSG ratio to that of diamide-induced
apoptosis. The results in Fig. 8
show that diamide causes a very rapid and significant decrease in cell
GSH at 15 min that recovered to control values by 1 h (Fig. 8A
). GSH levels at 2–4 h were higher than the baseline
values. In comparison, the GSH/GSSG ratio was also significantly
decreased at 15 min after diamide treatment, and remained suppressed
for 2 h while rapid recovery of cell GSH occurred within 1 h
(Fig. 8A
). Thereafter, the ratio returned to control status
by 4 h through 24 h. It is notable that during the first
2 h of severe imbalance in the GSH/GSSG status, apoptosis was low
and did not increase until 16 h and 24 h (Fig. 8B
), indicating that a sustained 2 h window of
disrupted redox balance was sufficient to elicit apoptosis and its
progression to cell death at 16 h and 24 h later.
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Diamide causes activation of caspase 3
To verify that the induction of GSH/GSSG imbalance by diamide is
temporally linked to activation of caspase 3, we examined the
expression of CPP32 and caspase-mediated cleavage of PARP. Figure 9A
shows that without diamide treatment, CPP32 was
undetectable in CaCo-2 cell extracts (lane 1), similar to the results
with LOOH (see Fig. 5A
, lane 1). Treatment of cells with 0.5
mM diamide resulted in an elevation of CPP32 expression at 15 min (lane
2) and a peak response at 30 min (lane 3). By 2 h the appearance
of procaspase 3 was decreased to near control levels (lanes 4 and 5).
These kinetic changes are consistent with a rapid expression the
proenzyme shortly after thiol oxidant challenge, followed by its
subsequent cleavage to active caspase 3, a temporal sequence similar to
that exhibited by exposing cells to LOOH (see Fig. 5A
).
Figure 9B
illustrates the kinetics of cleavage of PARP with
0.5 mM diamide treatment. The results show that diamide induces a
time-dependent cleavage of PARP to the 85 kDa and 29 kDa products, with
maximum cleavage occurring at 60 min after thiol oxidant exposure (lane
4). There was a decrease in the appearance of the 85 kDa and 29 kDa
products at 120 min (lane 5), which may reflect loss from secondary
proteolysis and/or protein underloading. Overall, the findings with
diamide and LOOH support a temporal sequence that involves
oxidant-induced early redox imbalance (15–30 min) that preceded
caspase 3 activation (60 min), PARP cleavage (60–120 min), and CaCo-2
cell apoptosis (24 h).
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Oxidative DNA damage is the result of LOOH-induced apoptosis
Recent studies by Esteve et al. have demonstrated an association
between apoptosis and oxidation of mitochondrial DNA (32)
.
To clarify whether oxidative DNA damage contributes to LOOH-induced
apoptosis of CaCo-2 cells, we determined the time course of the
formation of 8 oxoG after exposure to 10 µM LOOH. The results in
Fig. 10
show that LOOH treatment was associated with elevated concentrations of
8 oxoG at 48 h. Exposure to LOOH for 6 h to 24 h did not
significantly increase 8 oxoG formation (Fig. 10A
). In
contrast, significant increases in apoptosis were observed at 16 h
(15%), 24 h (20%), and 48 h (30%) after LOOH exposure
(Fig. 10B
). Notably, the increase in 8 oxoG levels at
48 h corresponded to maximal apoptosis; even though apoptosis was
elevated at 16 h and 24 h, DNA oxidation at these times was
not significant.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Shifting these control checkpoints in
the direction of reductant or oxidants during oxidative stress could
result in a cell that favors quiescence, proliferation, or death
(16
, 17)
. As pertains to a labile tissue like the
intestine, induction of oxidative shifts in the cellular redox status
can enhance cellular mitogenic or apoptotic responses
(17
18
19)
. Recent studies from our laboratory have
demonstrated that induction of mild redox imbalance at low
concentrations of LOOH (1 µM and 5 µM) promotes CaCo-2 cell
proliferation and cell cycle progression (Gotoh, Y. et al., unpublished
results), indicating a role for cellular redox status in modulating
intestinal cell proliferative activity. Studies from other laboratories
have demonstrated GSH depletion with apoptosis in response to different
apoptotic stimuli (32)
, indicating a link between
apoptotic cell death and altered cellular thiol status. Studies by
Garland et al. showed that apoptosis in murine interleukin 3 (IL-3)
-dependent homeopathic cells induced by IL-3 withdrawal lowered
cellular GSH but did not increase free radicals (34)
,
suggesting that cellular redox changes can modulate cell apoptosis
independent of oxidant generation.
The present study provides evidence to support our hypothesis that
LOOH-induced apoptosis in CaCo-2 cells is closely linked to
LOOH-induced disruption of normal cellular thiol/disulfide status. Our
results support a temporal link between LOOH induction of redox shifts,
activation of caspase 3, and initiation of apoptosis in CaCo-2 cells,
and are consistent with an early loss of redox balance that ultimately
leads to the apoptotic outcome. That LOOH-induced cell death is
characteristic of apoptosis is evidenced by flow cytometric analysis of
TUNEL-positive cells and the detection of distinct DNA fragments that
are consistent with oligonucleosome-length fragments of 180 bp on
agarose gels (31)
. It is notable that a physiologically
relevant LOOH concentration (10 µM) significantly perturbs the
cellular redox status in the absence of cell toxicity. The disruption
of cell GSH and GSSG occurred within 15–30 min of LOOH treatment,
indicating that loss of redox homeostasis contributes to the early
apoptotic responses to oxidant challenge. Moreover, this rapid kinetic
shift in the GSH/GSSG balance from acute LOOH exposure was associated
with activation of caspase 3, thus linking the loss of redox
homeostasis with initiation of the apoptotic cascade. Indeed,
LOOH-induced early loss of cellular redox balance may represent an
inciting event that governs LOOH-mediated apoptotic cell death in
intestinal cells. A similar link between GSH oxidation and apoptosis in
fibroblasts was observed with serum withdrawal (32)
.
Our kinetic studies of redox changes and apoptosis with diamide
treatment support the conclusion that rapid loss of GSH/GSSG
homeostasis is a critical upstream event in the apoptosis of CaCo-2
cells, as evidenced by the dissociation of diamide-induced redox
imbalance that occurred in 15 min from the first appearance of
apoptotic cells at 16 h later (Fig. 8)
. It is notable that whereas
treatment of cells with diamide resulted in a rapid disruption of
GSH/GSSG status similar to that induced by LOOH treatment, the
imbalance in redox status was sustained in LOOH- but not in
diamide-exposed cells. One reason that GSH was recovered in
diamide-treated cells may be because diamide oxidation of cell GSH
results in the elimination of the thiol oxidant, which allows for the
subsequent catalytic regeneration of GSH from GSSG by GSSG reductase.
In contrast, exposure of cells to LOOH will likely cause continuous
consumption of cellular GSH due to LOOH-mediated propagation of lipid
peroxidation and the generation of oxygen radicals. Yet both agents
elicited similar kinetics of early redox disruption that temporally
preceded caspase activation and cell apoptosis. This correspondence is
consistent with our suggestion that an initial disruption of redox
status is the critical step in oxidant-mediated cell apoptosis.
Moreover, our results suggest that the change in GSH/GSSG past 2 h
postoxidant challenge was not a major determinant of the apoptotic
outcome, given that the recovery of redox balance did not prevent
apoptosis in diamide-exposed cells. These results are consistent with
our overall hypothesis that sustained disruption of redox balance is
not necessary or critical to affect the ultimate apoptotic outcome.
Rather, there may exist an irreversible temporal checkpoint for redox
recovery such that failure to compensate for this imbalance will cause
apoptosis to continue to its final biological apoptotic end point
despite subsequent restoration of the cellular redox status. In our
studies, the threshold window for restoration of GSH/GSSG balance is
within the first 2 h after initiation of oxidant stress.
An interesting observation in the present study is the finding that
apoptosis in CaCo-2 cells was mediated predominantly by changes in the
GSH-to-GSSG ratio rather than by changes in levels of GSH per
se. This is evidenced by the finding that depletion of cellular
GSH alone using BSO without markedly altering the GSH/GSSG ratio did
not elicit apoptosis; rather, cell apoptosis was associated with
increased GSSG relative to GSH. Another important observation is the
finding that oxidative DNA damage did not contribute to the apoptotic
process; rather, that DNA oxidation occurs as a result of cells
undergoing apoptosis. A relationship between mitochondrial DNA damage
and apoptosis of the mammary gland was recently described
(32)
.
The mechanism by which LOOH induces apoptosis remains to be
identified. A reasonable sequelae of initiating events may involve
LOOH-induced loss of mitochondrial integrity, release of cytochrome
c, generation of mitochondrial oxy radicals that results in
the loss of cell redox balance, and the activation of caspase 3, a
mechanism that is consistent with mitochondrial redox signaling in
apoptosis (35
, 36)
. We are currently investigating whether
the mitochondrial redox signaling pathway is a major contributor to the
initiating event in LOOH-induced apoptosis in CaCo-2 epithelial cells.
Another mechanism that redox shift, resulting from LOOH metabolism, can
modulate apoptosis may be tied to its role in transcription factor
activation and binding to enhancer-promoter elements in DNA. Recent
studies have demonstrated that the binding of the nuclear factor 
B
(NFB) to DNA in HeLa cell extracts was increased by hydrogen
peroxide (37)
. Thus, a highly oxidized environment such as
occurs with increased oxy radical formation and loss of GSH homeostasis
could favor gene transcriptional activity. However, the kinetics of
LOOH-induced redox shift (by 30 min) and the activation of caspase 3
(by 1 h) suggest that a transcription-dependent mode of apoptotic
induction is unlikely in our model of oxidative stress. Moreover, the
results with diamide confirm that redox initiation of cell apoptosis
occurred early after oxidant exposure, which is consistent with a
transcription-independent initiating event. Notwithstanding, the above
considerations do suggest a potential role for redox modulation of
transcriptional activity of apoptotic genes, an avenue of research that
warrants further investigation. Regardless of the mechanism, our
present study provides evidence that redox perturbation induced by
subtoxic levels of lipid hydroperoxide leads to activation of caspase
3, oxidative DNA damage, and enhanced apoptosis. These results
underscore the importance of redox status in hydroperoxide-mediated
apoptosis and the potential contribution of this process to the genesis
of gut pathology.
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ACKNOWLEDGMENTS |
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Received for publication August 27, 1999.
Revision received February 16, 2000.
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REFERENCES |
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