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INTRODUCTION |
Solar UV irradiation reaching the surface of the earth, consisting
mainly of UVB (280-315 nm) and UVA (315-400 nm), presents a major
environmental challenge to the skin, contributing not only to
photo-aging but also to carcinogenesis (1-3). UVA has long been
considered less of a causative factor in skin carcinogenesis than UVB
due to its negligible absorption by DNA. However, the greater abundance
of UVA in solar UV irradiation and deeper penetration of UVA into the
actively dividing basal layer of the skin increases the relative
importance of UVA as compared with UVB (4). UVA has been shown to be a
risk factor for melanoma in fish (5) and could be also in humans
(6-8).
Fortunately, the carcinogenic effects of UV irradiation may be
decreased by apoptosis, programmed cell death, which eliminates DNA-damaged or potentially mutated cells. Much more is known about UVB-induced apoptosis than about apoptosis induced by UVA in both normal keratinocytes and immortalized keratinocytes (9-19). However, Godar and co-workers (20-22) have reported that UVA-induced apoptosis in lymphoma and Jurkat cells involves mechanisms that differ from those
seen with UVB. Oxidative stress is presently considered to be involved
in UVA-induced apoptosis (3). In support of this involvement, a
protective role is played by tea polyphenols against both cytotoxicity
and apoptosis induced by UVA in rat keratinocytes; this protection
correlates well with the ability of the polyphenols to quench reactive
oxygen species (23). Nevertheless, the mechanisms involved in
UVA-induced apoptosis of human keratinocytes are still poorly understood.
There is evidence that protection of human skin cells against a wide
range of solar UV radiation damage, including UVA, involves endogenous
glutathione (24). This thiol is involved in many biological processes,
including the regulation of gene expression, apoptosis, and membrane
transport (25-27). Glutathione is considered to be the most prevalent
and most important intracellular non-protein thiol-disulfide redox
buffer in mammalian cells (25-27). The higher glutathione content in
immortalized HaCaT cells is expected to confer resistance to UVA
irradiation as compared with normal keratinocytes (28).
In apoptosis, the role of glutathione is controversial and dependent on
cell types and pro-apoptotic stimuli. (i) High intracellular reduced
GSH levels have been found to prevent Fas-induced cell apoptosis in the
Fas-resistant variant CEM2D1R (29), whereas decreased GSH levels
enhance Fas-induced apoptosis in the Fas-resistant variant CEM2D1R and
the human hepatoma cell line HepG (2, 29, 30). (ii) Low intracellular
GSH levels have also been shown to prevent apoptosis by compromising
caspase activation in mouse hepatocytes (31, 32). Depletion of
intracellular GSH prevented CD95-triggered apoptosis upstream of
caspase-8 activation in T and B cells (33). Furthermore, cells
undergoing apoptosis also appear to export GSH into the extracellular
space (34-37). However, neither the mechanism involved in the
transport of GSH nor the functional benefit of this process is known.
In this study we have found that in immortalized HaCaT cells, UVA
irradiation induces the active efflux of GSH. We have surveyed the
mechanisms involved in this efflux, focusing on the possible roles of
MRP1 transporter protein. We
also found that GSH export is closely associated with UVA-induced apoptosis.
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EXPERIMENTAL PROCEDURES |
Materials--
Reduced GSH, GSSG, verapamil (VP), cyclosporin A
(CsA), 5-sulfosalicylic acid glutathione reductase, and buthionine
sulfoximine (BSO) were purchased from Sigma.
5,5'-Dithiobis(2-nitrobenzoic acid) (DTNB), 2-vinylpyridine, EDTA,
glucose, NADPH, and triethanolamine were purchased from Aldrich.
Cell Culture--
The spontaneously immortalized human
keratinocyte cell line HaCaT (38), obtained from Prof. N. Fusenig
(German Cancer Research Center, Heidelberg, Germany), was maintained in
monolayer culture in 95% air, 5% CO2 at 37 °C in
Dulbecco's modified Eagle's medium supplemented with 10% fetal
bovine serum, 31 µg/ml penicillin, and 50 µg/ml streptomycin. For
experiments, HaCaT keratinocytes were grown in 12-well plates or
plastic Petri dishes (100 mm) for 24-48 h. 4 h prior to UVA
treatment, subconfluent cells were given fresh Dulbecco's modified
Eagle's medium containing 1% fetal bovine serum. For GSH depletion,
BSO (50 µM) was added to the culture medium, and the
cells were incubated for 18 h prior to UVA exposure (39); this
treatment had no effect on cell viability.
UVA Treatment--
The medium was removed, and cells were washed
once with sterile PBS (PBS-CMF, calcium/magnesium-free). After the
addition of sterile PBS or PBS containing 10 mM of glucose,
the cells were irradiated with fluorescent lamps (Houvalite F20T12BL-HO
PUVA, National Biological Corp., Twinsburg, OH) with the dish lid on. The UVA dose was monitored with a Goldilux UV meter equipped with a UVA
detector (Oriel Instruments, Stratford, CT). Control samples were kept
in the dark under the same conditions. After treatment, the supernatant
was removed or collected as indicated, and the cells were washed with
PBS. For the apoptosis assay, fresh medium containing 1% fetal bovine
serum was added after exposure, and the cells were incubated at
37 °C. At predetermined time points, attached and floating cells
were harvested and subjected to apoptosis analysis. In selected
experiments, cells were pretreated with GSH or inhibitors for multidrug
resistance-associated protein (MRP) at 37 °C for 15 min prior
to irradiation.
Determination of Intracellular and Extracellular
Glutathione--
GSH and GSSG were measured using a modified method
for glutathione determination in microtiter plates (40, 41). Briefly, after removal of the supernatant, cells in 12-well plates were washed
with PBS and then treated with 500 µl of 10 mM
hydrochloric acid and stored at
20 °C until analyzed. Cells were
scraped and sonicated three times for 5 s and then centrifuged at
10,000 × g for 10 min at 4 °C. Aliquots (350 µl)
of the supernatant were collected, of which 50 µl were transferred to
separate microtubes for BCA protein assay (Pierce). To the remaining
aliquot (300 µl), 300 µl of 5% 5-sulfosalicylic acid was added,
and the mixture was vortexed and then kept on ice for 5 min to
precipitate protein. After centrifugation at 10,000 × g for 10 min at 4 °C, 500 µl of supernatant were
collected, neutralized by the addition of 4 M
triethanolamine solution, and divided into 2 aliquots for measurement
of GSSG and total glutathione. To conjugate GSH, 2-vinylpyridine was
added to one of the aliquots to a final concentration of 2% (v/v).
The microtiter plate was prepared by pipetting 50 µl of standards or
samples per well. Immediately, 100 µl of freshly prepared assay mix
(0.49 ml of 6 mM DTNB, 3.75 ml of 1 mM NADPH,
8.15 ml of phosphate/EDTA buffer, and 20 units of glutathione
reductase) was pipetted into each well. The plate was placed into the
microplate reader (Tecan US SPECTRAFluor Plus, Research Triangle Park,
NC) and the absorption at 405 nm monitored. The GSH or GSSG levels were
expressed as nmol/mg protein or ratio as compared with respective controls without UVA treatment. For extracellular GSH and GSSG, cell
supernatants were treated similarly, with or without 2-vinylpyridine followed by microtiter detection using the DTNB/NADPH/glutathione reductase assay mix.
DNA Fragmentation--
The pattern of DNA cleavage was analyzed
by agarose gel electrophoresis. Briefly, cell pellets were resuspended
in lysis buffer (5 mM Tris-HCl, pH 8.0; 20 mM
EDTA; 0.5% Triton X-100) and incubated on ice at 4 °C overnight.
After incubation at 56 °C for 1 h with RNase A (100 µg/ml)
and then 1 h with proteinase K (200 µg/ml), the cell lysate was
extracted with phenol/chloroform/isopropyl alcohol (25:24:1, v/v). DNA
was precipitated with ethanol and subsequently washed with 70%
ethanol. DNA samples, dissolved in 1× TE buffer, were separated by
horizontal electrophoresis on 1.5 or 1.8% agarose gels, stained with
ethidium bromide, and visualized under UV light.
Caspase Activity--
Caspases were assayed using ApoAlert
Caspase Fluorescent Assay Kits (Clontech, Palo
Alto, CA). Briefly, cells were extracted in lysis buffer, and the cell
lysate was incubated for 1 h at 37 °C with assay buffer
containing one of the following fluorescent caspase substrates:
Ac-DEVD-AFC for caspase-3, Ac-IETD-AFC for caspase-8, and Ac-LEHD-AFC
for caspase-9. The fluorometric detection of cleaved AFC product was
performed on a plate reader (excitation 400 nm and emission 505 nm).
For preparation of the AFC calibration curve, 80 µM free
AFC was diluted in the caspase assay buffer without substrate to give
0.5, 1, 2, and 4 µM of free AFC. The results were
expressed as ratio of the treated samples to respective control samples.
Lactate Dehydrogenase (LDH) Release--
LDH, a stable cytosolic
enzyme that is released upon the increase in plasma membrane
permeability, is determined by CytoTox96 non-radioactive cytotoxicity
assay kit (Promega, Madison, WI). Briefly, after treatment, 50 µl of
medium was taken at different time points to the enzymatic assay plate.
After all samples were taken, 50 µl of substrate mix was added to
each well of the enzymatic assay plate, and the plate was incubated at
room temperature for 30 min in the dark. The reaction was stopped by
adding 50 µl of stop solution, and LDH was determined by recording
the absorbance at 492 nm by a plate reader. The maximum LDH release
control was done by adding lysis solution to the control cells. The
results were expressed as LDH released (%) = (treated
control)/(maximum
control) × 100.
Flow Cytometry--
Annexin V staining was used to determine the
translocation of phosphatidylserine (PS) in UVA-induced apoptosis.
After incubation, control or treated cells including attached and
floating cells were harvested and collected by centrifugation at
300 × g for 5 min at room temperature. Cells were
washed with cold PBS and stained with TACSTM annexin V kits
according to the manufacturer's instructions (Trevigen, Gaithersburg,
MD). Cells positive for annexin V-fluorescein isothiocyanate were
quantified by flow cytometry using a BD FACSort (BD Biosciences).
Statistics--
Data are presented as mean ± S.E. of three
to six experiments. The Student's t test was used for
comparisons between experimental groups (n = 3-6). A
value of p < 0.05 was considered statistically significant.
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RESULTS |
UVA-Induced GSH Efflux in HaCaT Cells--
In order to determine
the effect of UVA on the efflux of GSH, we irradiated subconfluent
HaCaT cells in PBS with UVA, with and without glucose. We found that in
the presence of glucose to provide energy, UVA (25 J/cm2)
induced significant efflux of GSH (Fig.
1).

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Fig. 1.
UVA induced GSH efflux in the presence of
glucose in HaCaT cells. Intracellular (A) and
extracellular (B) GSH, GSSG, and total glutathione levels
immediately after the HaCaT cells were switched from normal culture
medium to PBS or 10 mM glucose PBS and then sham-irradiated
or UVA-irradiated (25 J/cm2). Results are the mean ± S.E. of three to six experiments (**, p < 0.01 as
compared with dark control samples). C, the cells were
treated as in A and B with different doses of
UVA. The extracellular glutathione ratios of UVA-irradiated samples to
their respective control samples were determined. D, cells
were irradiated with 25 J/cm2 UVA in the presence of
different glucose concentrations. The extracellular glutathione ratios
of UVA-irradiated samples to their respective control samples were
determined. The GSH efflux depended on the concentration of glucose.
Cells were seeded in 12-well plates and used after 48 h of
culture. After washing with PBS once, medium was replaced by an
equivalent quantity of PBS or glucose PBS, and the plates were
irradiated with UVA. Control samples were kept in the dark. For
intracellular glutathione determination, cells were harvested after
treatment and prepared for total glutathione and GSSG assay by DTNB
method as described under "Experimental Procedures." The
supernatants were collected for extracellular total glutathione and
GSSG assay by the same method.
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The total glutathione level in HaCaT cells in normal culture is 62.6 nmol/mg protein, of which more than 98% is in the reduced form (GSH).
As compared with normal cells cultured in medium, intracellular total
glutathione, GSH + GSSG, of cells incubated in the dark decreased to
41.9 and 40.6 nmol/mg protein for PBS and glucose PBS solutions,
respectively (Fig. 1A). GSH levels also decreased, whereas
levels of the oxidized form, GSSG, increased ~3-fold. When HaCaT
cells were irradiated with UVA at a dose of 25 J/cm2 in the
presence of glucose (10 mM), intracellular total
glutathione levels and GSH decreased significantly from dark controls,
whereas GSSG levels remained unchanged (Fig. 1A). Thus the
decrease in total glutathione appears to be due to a reduction in GSH.
To determine whether GSH was exported into the extracellular medium,
supernatants were collected, and GSH, GSSG, and total glutathione
levels were determined (Fig. 1B). In the dark, the presence
of glucose had no effect on the levels of extracellular GSH, GSSG, or
total glutathione. When the sum of the intracellular and extracellular
total glutathione was compared with intracellular total glutathione in
normal cultured cells, the loss of intracellular total glutathione was
recovered completely in the supernatant. Efflux of GSH caused a
decrease in intracellular GSH. In the absence of glucose, however,
there was no GSH efflux, and the intracellular GSH depletion was due to
the oxidation to GSSG (Fig. 1, A and B).
When HaCaT cells were irradiated with different doses of UVA (Fig.
1C), GSH efflux increased with the UVA dose. There was no
GSH efflux below 10 J/cm2, but GSH efflux increased up to
25 J/cm2 and remained unchanged. Interestingly, consistent
with Fig. 1, A and B, no GSH efflux was observed
under different doses of UVA irradiation in the absence of glucose. To
confirm the requirement of glucose, GSH efflux was determined in the
presence of different concentrations of glucose (Fig. 1D).
Under UVA irradiation (25 J/cm2), GSH was exported in a
glucose concentration-dependent manner. These results
suggest that glucose is required for GSH efflux induced by UVA irradiation.
Effect of Exogenous GSH Addition and GSH Depletion on GSH
Efflux--
To determine whether the efflux of GSH under UVA
irradiation is influenced by GSH levels, we used exogenous GSH to
increase extracellular GSH levels or BSO, a known GSH-depleting agent, to decrease intracellular GSH. Each agent was added and incubated with
cells prior to UVA irradiation.
In the dark, exogenous GSH (1 mM) incubated with cells for
15 min at 37 °C did not affect intracellular GSH or GSSG even in the
presence of glucose (Fig. 2A).
As observed previously, when cells were irradiated with 25 J/cm2 of UVA in the presence of glucose, GSH was exported
into the extracellular medium (see above). However, the addition of
exogenous GSH reduced the loss in intracellular GSH and thus the
UVA-induced efflux significantly. The GSSG level did not change
significantly with or without exogenous GSH (Fig. 2A).

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Fig. 2.
Effect of the addition of GSH and GSH
depletion by BSO on UVA-induced GSH efflux. Intracellular
(A) and extracellular (B) GSH, GSSG, and total
glutathione levels when the HaCaT cells were pretreated with GSH (1 mM) or BSO and then kept in the dark or UVA-irradiated (30 J/cm2). For GSH treatment, after 48 h of culture cells
were washed once with PBS, after which glucose PBS (10 mM)
containing 1 mM of GSH was added, and cells were incubated
at 37 °C for 15 min prior to irradiation. After irradiation, cells
were washed twice with PBS to eliminate the effect of exogenous GSH on
the determination of intracellular GSH levels. For BSO treatment, BSO
(50 µM) was added 18 h before irradiation; cells
were then washed with PBS twice, after which glucose PBS was added
prior to irradiation. For intracellular glutathione determination,
cells were harvested after irradiation and prepared for intracellular
GSH and GSSG assay by the DTNB method. The supernatants were collected
for extracellular GSH and GSSG assay by the same method. Results are
the mean ± S.E. of three to six experiments. (*,
p < 0.05; **, p < 0.01 as compared
with the UVA-irradiated samples without GSH or BSO).
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When BSO (50 µM) was added to the cell culture 18 h
prior to UVA exposure, intracellular total glutathione decreased
dramatically; no cytotoxicity was detected (data not shown). As was the
case with non-BSO-pretreated cells, the UVA-induced decrease in total intracellular glutathione in BSO-pretreated cells was recovered from
the extracellular medium (Fig. 2B). Although GSH efflux was induced by UVA irradiation after GSH depletion by BSO incubation, the
efflux process slowed down due to the dramatic decrease in intracellular GSH.
Effect of MRP1 Inhibitors on UVA-induced GSH Efflux--
To
identify the possible transport carrier involved in UVA-induced GSH
efflux, inhibitors for multidrug resistance-associated protein 1 (MRP1)
were preincubated with cells prior to irradiation. In the dark, VP (10 or 20 µM) and CsA (5 or 10 µM) caused no
significant change (data not shown). When cells pretreated with
verapamil or CsA were irradiated with UVA, UVA in the presence of
either inhibitor caused less of a GSH efflux as compared with UVA alone (Fig. 3A). Higher
concentrations of CsA or VP exhibited more inhibition on GSH efflux.
These results indicated that verapamil and CsA both inhibited the
efflux of GSH, with CsA being somewhat more effective. The much lower
inhibition by verapamil could be due to the fact that verapamil
inhibits MRP activity non-competitively and may exhibit lower
inhibition on the binding of GSH with MRP1, as compared with
cyclosporin A, a competitive inhibitor of MRP1-substrate binding.

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Fig. 3.
CsA and VP inhibited UVA-induced GSH efflux
in HaCaT cells. A, cells were incubated with CsA (5 or
10 µM) or VP (10 or 20 µM) at 37 °C for
15 min and then irradiated with UVA (25 J/cm2) in the
presence of 10 mM glucose. Immediately after irradiation,
supernatants were taken to measure the extracellular glutathione. The
results are expressed as the ratio of treated levels to respective
control levels. B, after UVA irradiation (25 J/cm2), cells were given fresh medium. At predetermined
time points, medium was taken to determine the extracellular
glutathione levels. Results are the mean ± S.E. of three to six
experiments. (*, p < 0.05; **, p < 0.01 as compared with the UVA-irradiated samples without CsA or
VP).
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However, after irradiation, UVA did not cause further GSH efflux as
compared with the control samples (Fig. 3B). Neither
verapamil nor cyclosporin A affected the GSH efflux. This suggests that UVA-induced GSH efflux occurs during the irradiation but not after irradiation.
Role of GSH Efflux in UVA-induced Apoptosis--
To evaluate the
physiological role of UVA-induced GSH efflux, we examined the influence
of GSH efflux inhibition on UVA-induced apoptosis. HaCaT cells were
first treated with or without cyclosporin A, verapamil, or GSH and then
subjected to UVA irradiation. DNA fragmentation was monitored by
electrophoresis; the activities of caspases-3, -8, and -9 were measured
with their respective fluorescent substrates; the plasma membrane
permeability was determined by LDH release; and the externalization of
PS on the cell surface was monitored by flow cytometry in combination
with annexin V staining.
As shown in Fig. 4, A and
B, UVA irradiation caused a dose- and
time-dependent increase in DNA fragmentation in HaCaT
cells. However, UVA-induced DNA fragmentation was not markedly
inhibited by prior treatment of HaCaT cells with cyclosporin A,
verapamil, or GSH (Fig. 4C). UVA irradiation activated
caspases-3, -8, and -9 in a time-dependent manner (Fig.
5A). After UVA irradiation (25 J/cm2), caspases-3, -8, and -9 were activated as early as
30 min. However, pretreatment with cyclosporin A, verapamil, or GSH did
not affect the activation of these caspases at 1 (Fig. 5B)
or 6 h (Fig. 5C). Furthermore, the pretreatment of the
cells with Ac-DEVD-CHO, the caspase-3 inhibitor, had no effect on
UVA-induced GSH efflux (Fig. 5D). There was no caspase-3
activation (data not shown) while dramatic GSH efflux occurred (Fig. 1)
immediately after UVA exposure, suggesting that GSH efflux induced by
UVA precedes the caspase-3 activation.

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Fig. 4.
Inhibition of GSH efflux has no effect on
UVA-caused DNA fragmentation. A, cells were irradiated
with different doses of UVA (0, 10, 20, 25, 30, and 40 J/cm2) in the presence of 10 mM glucose and
then incubated for 15 h. DNA was extracted and separated on 1.5%
agarose gel. B, cells were irradiated with 25 J/cm2 UVA and incubated for different times (3, 6, 12, 15, 18, and 24 h). Extracted DNA was subjected to 1.5% agarose gel
electrophoresis. C, cells were preincubated with CsA (5 or
10 µM) or VP (10 or 20 µM) at 37 °C for
15 min and them irradiated with UVA (25 J/cm2) in the
presence of 10 mM glucose. After incubation for 15 h,
DNA from treated and untreated samples was separated on 1.8% agarose
gel. Cells were seeded in 100-mm Petri dishes and used at a 50-70
confluence. Results are representative of three independent
experiments.
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Fig. 5.
Inhibition of GSH efflux has no effect on the
activation of caspases-3, -8, and -9 by UVA. A, after
UVA exposure (25 J/cm2), the cells were incubated for
different times. The caspase activity was determined in cytosolic
extracts prepared at the time point given. B, in a parallel
experiment, the cells were pretreated with CsA (10 µM),
VP (20 µM), or GSH (1 mM) and then irradiated
with UVA (25 J/cm2). The caspase activity was determined
1 h after UVA treatment. C, the cells were treated the
same as in B, and the caspase activity was determined 6 h after UVA irradiation. D, the cells were pretreated with
caspase-3 inhibitor, Ac-DEVD-CHO (10 and 20 µM). After
UVA irradiation (25 J/cm2), the supernatants were taken,
and glutathione levels were determined. Results are the mean ± S.E. of four to six experiments.
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UVA also caused a gradual increase in LDH release in HaCaT cells in a
dose- and time-dependent manner (Fig.
6A). The presence of
cyclosporin A (2.5, 5, or 10 µM) (Fig. 6B),
verapamil (5, 10, or 20 µM) (Fig. 6C), or GSH
(0.2 or 1 mM) (Fig. 6D) inhibited the LDH
release. It is evident that GSH is more efficient in inhibiting UVA-caused LDH release than CsA or VP. These results suggest that GSH
efflux induced by UVA is involved in the increased in plasma membrane
permeability. Cell lysis especially occurs in the late stage of
apoptosis when macrophages are absent.

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Fig. 6.
Inhibition of GSH efflux decreased the LDH
release induced by UVA. A, the cells were irradiated
with different doses of UVA (10, 20, 25, 30, and 40 J/cm2)
as indicated. Supernatant was taken at different time points to
determine the LDH release. In a parallel experiment, the cells were
pretreated with different concentrations of CsA (B), VP
(C), or GSH (D) as indicated and then irradiated
with UVA (25 J/cm2). At different time points given,
supernatant was taken to determine the LDH release. Results are the
mean ± S.E. of four to six experiments. (*, p < 0.05 as compared with the UVA-irradiated samples without CsA, VP, or
GSH).
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In order to test whether GSH efflux is involved in the translocation of
PS, a membrane lipid rearrangement occurring in the early or
intermediate stage of apoptosis and the triggering event in the
recognition of apoptotic cells by the scavenger receptors of
macrophages (42), annexin V was used to identify PS exposed on the
external membrane surface following rearrangement of the lipid
bilayer. As shown in Fig. 7, UVA induced
a dramatic increase in PS translocation as early as 4 h after
exposure. However, the pretreatment with CsA (10 µM), VP
(20 µM), or GSH (1 mM) significantly inhibited PS translocation at both 4 and 18 h after UVA
irradiation. These findings suggest that GSH efflux contributes to
UVA-induced PS translocation in HaCaT cells.

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Fig. 7.
Inhibition of GSH efflux decreased the
translocation of PS in UVA-induced apoptosis. Cells were
preincubated with CsA (10 µM), VP (20 µM),
or GSH (1 mM) at 37 °C for 15 min and then irradiated
with UVA (25 J/cm2). After incubation for 4 or 18 h,
the cells were harvested to determine the binding of annexin V to the
externalized PS on the cell surface (55, 60). Results are the mean ± S.E. of four to six experiments. (**, p < 0.01; *,
p < 0.05 as compared with UVA-irradiated samples
without CsA, VP, or GSH at 4 and 18 h after UVA exposure,
respectively).
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It is noteworthy that inhibition of MRP1 is not the only effect of
verapamil and cyclosporin A. Verapamil is known to be a calcium channel
blocker and protect methoxyacetic acid-induced spermatocyte apoptosis
in cultured rat seminiferous tubules (43). Cyclosporin A increased
K+-induced calcium influx (44) and mitochondria calcium
storage (45). Although the presence of verapamil or cyclosporin A might influence calcium homeostasis, no effect on DNA fragmentation or
caspase activation, the two hallmarks for apoptosis, was observed in
our study. This is the same with the addition of exogenous GSH. On the
other hand, similar effect was observed by the presence of verapamil,
cyclosporin A with exogenous GSH, which all inhibited GSH efflux and PS
translocation. Therefore, we proposed that the effect of CsA and
verapamil on UVA-triggered apoptosis was at least partially due to
the inhibition of MRP1.
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DISCUSSION |
Recent studies have revealed that several types of cells
undergoing apoptosis induced by pro-apoptotic stimuli, such as
diphenyleneiodonium, puromycin, ricin, and anti-Fas/APO-1 antibody,
appeared to release GSH rapidly and selectively into the extracellular
space (34-37). However, neither the mechanisms involved in the
transport of GSH nor the biological functional significance of this
cellular GSH release is known. Prior to this study the involvement of
GSH efflux in apoptosis of epidermal keratinocytes induced by UVA
irradiation, an important environmental stress factor, was also unknown.
In the present study, we have tested the putative mechanisms involved
in GSH efflux and apoptosis of HaCaT cells induced by UVA. We have
shown that exposure of HaCaT cells to UVA irradiation increased the
transport of reduced GSH into the extracellular medium and that
UVA-induced apoptosis was closely associated with this efflux.
It is evident that UVA irradiation caused significant glutathione
efflux in HaCaT cells considering the decreased intracellular total
glutathione and increased extracellular total glutathione, provided
that an energy source (glucose) was present. For cells kept in the
dark, the efflux of intracellular GSH was much smaller, probably
resulting from cystine withdrawal when the cells were switched from
normal culture medium to cystine-free PBS or glucose PBS solution (35,
46).
We found that UVA-induced GSH efflux depended on the presence of
glucose. Thus the transport of GSH into the extracellular space is an
active process and energy-dependent (35, 47) and is not due
to passive leakage as a result of the loss of membrane integrity.
Immediately after UVA irradiation, only about 2% of cells were
propidium iodide-positive and had lost their membrane integrity
(data not shown). It seems unlikely that these damaged cells could be
responsible for an increase in GSH efflux of almost 70%. Without
glucose, UVA irradiation did not induce a greater GSH efflux than in
dark controls, suggesting that GSH efflux under these conditions is due
to cystine withdrawal following substitution of PBS for medium.
Evidently the presence of glucose not only provides the energy for
active transport of GSH but also maintains lower intracellular GSSG
levels and higher GSH levels under UVA stress by providing a source of
NADPH that is essential for the recycling of GSSG to GSH (48, 49).
In previous studies, under UVA irradiation in the absence of glucose,
the oxidation of both intracellular and extracellular GSH to GSSG was
found to be the predominant reaction for glutathione, causing the
depletion of intracellular GSH (24, 50). No significant GSH efflux was
observed, as was the case in our studies. The efflux of GSH was
inhibited by exogenous addition of GSH and slowed down by GSH depletion
(80%) with BSO. This suggests that the transport process is affected
by the concentration gradient of GSH across the cell membrane.
As far as the mechanisms and carriers involved in GSH transport are
concerned, it is known that the multidrug resistance-associated proteins, MRP1 and MRP2, are responsible for GSH efflux in the liver
(47, 51, 52). Because the expression of MRP2 is restricted to liver,
kidney, and gut, whereas MRP1 is found in many tissues (53), it is
reasonable to suppose that in HaCaT keratinocytes, MRP1 may be the
transporter protein that is activated by UVA irradiation to cause GSH
efflux. Furthermore, the rapid efflux of GSH during UVA irradiation
(within 2 h) suggests that this active transport process may be
due to the activation of pre-existing transporter proteins by UVA in
the presence of the energy provider glucose, rather than the induction
of transporter proteins by UVA irradiation or oxidative stress (54).
Verapamil and CsA, inhibitors for MRP1, prevent the efflux of GSH
induced by UVA irradiation, which implies the participation of MRP1 in
the transport process of GSH in HaCaT cells.
The rapid and active efflux of GSH induced by UVA irradiation in HaCaT
cells could be an important biological response of keratinocytes to
UVA-induced stress. Although little is known about the biological
significance of GSH efflux in keratinocytes, we have found that
apoptosis of HaCaT cells that occurred after GSH efflux is closely
related to the specific GSH export process. GSH efflux has no effect on
caspase activation and DNA fragmentation in our study. However, the
increases in plasma membrane permeability and PS translocation during
apoptosis of HaCaT cells are associated with GSH efflux. Inhibition of
GSH efflux by the MRP1 inhibitors verapamil and CsA also prevents the
loss of plasma membrane integrity and PS externalization. The presence
of exogenous GSH has a similar effect to CsA and verapamil. Previous
studies (56, 57) have shown that PS translocation is independent of
both nuclear activity (55) and caspase activation. It seems most likely
that in HaCaT cells GSH efflux is involved in membrane rearrangement
rather than caspase activation or DNA fragmentation in UVA-induced
apoptosis. As a matter of fact, in apoptosis of Jurkat lymphocytes
induced by anti-Fas/APO-1, GSH efflux was observed during apoptosis,
but inhibition of this process did not affect DNA fragmentation (35). Inhibition of GSH efflux has been shown to protect cells from apoptosis
induced by puromycin in HepG2 cells but not in U937 cells by inhibiting
DNA fragmentation (34). As in the case of puromycin-treated HepG2 cells
(34), UVA-induced GSH efflux in HaCaT cells precedes DNA fragmentation;
however, DNA fragmentation is not affected by inhibition of GSH efflux.
It appears that cells stimulated by UVA to undergo apoptosis get rid of
their GSH during the irradiation in order to optimize the functioning
of the overall process. GSH efflux preceding apoptosis provides a novel
mechanism to deplete GSH in the cells rather than oxidizing GSH to
GSSG, which enhances the oxidative tonus without intervention of
reactive oxygen species (34, 35). The increased oxidative stress has
been shown to result in the selective phosphatidylserine oxidation that
precedes phosphatidylserine externalization in oxidant-induced
apoptosis (58, 59). Because externalized phosphatidylserine
serves as an important signal for targeting recognition and elimination
of apoptotic cells by macrophages (42), the earlier phosphatidylserine
translocation increased the macrophage recognition and phagocytosis of
the apoptotic cells in vivo and thus minimized their
potential to invoke inflammation or form neoplastic transformation.
These changes may be important for skin health and the prevention of
UVA-induced skin cancer.