Diethylstilbestrol Induces Rat Spermatogenic Cell Apoptosis
in Vivo through Increased Expression of Spermatogenic
Cell Fas/FasL System*
Radhika
Nair and
Chandrima
Shaha
From the National Institute of Immunology, Aruna Asaf Ali Road,
New Delhi, India 110067
Received for publication, September 11, 2002, and in revised form, November 6, 2002
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ABSTRACT |
The significant role that estrogens play
in spermatogenesis has opened up an exciting area of research in male
reproductive biology. The realization that estrogens are essential for
proper maintenance of spermatogenesis, as well as growing
evidence pointing to the deleterious effects of estrogen-like chemicals
on male reproductive health, has made it imperative to dissect the role estrogens play in the male. Using a model estrogen, diethylstilbestrol (DES), to induce spermatogenic cell apoptosis in vivo in
the male rat, we provide a new insight into an
estrogen-dependent regulation of the Fas-FasL system
specifically in spermatogenic cells. We show a distinct increase in
Fas-FasL expression in spermatogenic cells upon exposure to
diethylstilbestrol. This increase is confined to the spermatid
population, which correlates with increased apoptosis seen in the
haploid cells. Testosterone supplementation is able to prevent
DES-induced Fas-FasL up-regulation and apoptosis in the
spermatogenic cells. DES-induced germ cell apoptosis does not occur in
Fas-deficient lpr mice. One other important finding is that
spermatogenic cells are type II cells, as the increase in Fas-FasL
expression in the spermatogenic cells is followed by the cleavage of
caspase-8 to its active form, following which Bax translocates to the
mitochondria and precipitates the release of cytochrome c
that is accompanied by a drop in mitochondrial potential. Subsequent to
this, activation of caspase-9 occurs that in turn activates caspase-3
leading to the cleavage of poly(ADP-ribose) polymerase. Taken together,
the data indicate that estrogen-like chemicals can precipitate
apoptotic death in spermatogenic cells by increasing the expression of
spermatogenic cell Fas-FasL, thus initiating apoptosis in the same
lineage of cells through the activation of the apoptotic pathway chosen
by type II cells.
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INTRODUCTION |
The study of the role of estrogens in spermatogenesis has
attracted significant interest because of the increasing awareness that
estrogen is essential for spermatogenesis (1, 2), spermatogenic cells
express estrogen receptors
(ER)1 (1-3), and
estrogen-like chemicals present in the environment adversely affect
male reproductive health (4). Such chemicals have the ability to affect
gene expression and cellular function by binding to the hormone
receptors (5) and have been implicated in the declining trend of male
fertility and an increase in testicular cancers (4). Effects of
endocrine disruptors (ED), a term used to describe agents that mimic
hormones have been particularly well documented in wildlife populations
(6). In the male animal, one of the most susceptible pathways that can
be disrupted by EDs is the hypothalamic-pituitary-gonadal axis. This
axis regulates spermatogenesis by controlling the circulating levels of
luteinizing hormone and follicle-stimulating hormone through the
feedback regulation of steroid hormones (4), and this feedback loop can
be intercepted by EDs binding to ERs in the pituitary. Direct interference with spermatogenic cells is also possible because these
cells are known to express ERs (7). Diethylstilbestrol (DES) is a
stilbene estrogen that can bind to ERs in the pituitary and has been
widely used as a model estrogen to study changes in male reproductive
function in response to estrogens (8, 9). DES can mimic estrogen action
by interfering with the functioning of the pituitary-gonadal axis,
leading to the suppression of testosterone levels that result in
increased spermatogenic cell apoptosis (4, 10). Therefore, DES-induced
spermatogenic cell death is a suitable model to study the pathways
involved in estrogen-induced spermatogenic cell apoptosis.
Regular apoptosis of spermatogenic cells is required to maintain proper
testicular homeostasis, although increased cell death can result in
defective spermatogenesis leading to infertility (11). As in other
cells, male spermatogenic cells respond to external signals and to
their internal milieu by activating intracellular signaling pathways
that ultimately determine their fate. Signals that induce programmed
cell death are known to initiate apoptotic pathways in spermatogenic
cells involving members of the Bcl-2 family and also the Fas-FasL
system (12). In FasL-induced spermatogenic cell death, it is generally
accepted that FasL from Sertoli cells kill the spermatogenic cells by
engaging the Fas receptors present on them. The Fas system of
regulation in the testis is operative under various conditions of
stress like ethanol injury (13), heat exposure (14), and cryptorchidism
(15), where increases in Sertoli cell FasL expression have been
described but no spermatogenic cell FasL expression was reported.
However, recently, it has been shown that FasL is expressed in
spermatogenic cells (16). To our knowledge, no study has so far shown
the up-regulation of the Fas-FasL system specifically in the
spermatogenic cells as a means of initiating cell death in response to
any stimuli. Although the Fas-FasL system is reportedly important for
spermatogenic cell apoptosis, a functional Bax protein is also known to
be crucial, as expression of this protein helps initiate massive
spermatogenic cell apoptosis at a critical time during testicular
maturation (17).
Even though it is well established that estrogens induce spermatogenic
cell apoptosis, the dissection of complete pathways leading to
apoptotic death because of exposure to estrogens have not been worked
out. With increasing data accumulating on the role of estrogens in
spermatogenesis and the detrimental effects of environmental estrogens
on the same process (4), it has become essential to investigate the
mechanism of estrogen-induced apoptosis to understand spermatogenetic
disorders. Most of the studies addressing cellular apoptosis in
the testis using different model systems have used total testicular
tissue and not isolated spermatogenic cells to dissect pathways leading
to cell death (13-15). Failure to use isolated spermatogenic cells
might provide a completely different view, as the testis is a
heterogeneous organ composed of multiple cell types. To circumvent the
possibility of introducing variables contributed by other cell types,
we have used a purified spermatogenic cell population that is devoid of mature testicular sperm. This study projects a new possibility that
under conditions of estrogen exposure in vivo, increased expression of the spermatogenic cell Fas-FasL system may be responsible for initiating apoptotic death in the same lineage of cells by inducing
translocation of Bax from the cytosol to the mitochondria, followed by
the release of cytochrome c accompanied by a loss of
mitochondrial membrane potential (
m), leading to the
activation of caspase-9 and -3.
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EXPERIMENTAL PROCEDURES |
Materials--
The Apoptosis Detection System was procured from
Promega (Madison, WI). Bicinchoninic protein assay reagents A & B were
purchased from Pierce. Antibodies and secondary antibodies were
from Santa Cruz Biotechnology Inc. (Santa Cruz, CA) or PharMingen (San
Diego, CA) and Jackson Immunoresearch (West Grove, PA). Caspase assay kit was from Bio-Rad. For color development, VIP peroxidase substrate kit for Western blots and diaminobenzidine kit for immunohistochemical staining were procured from Vector Laboratories Inc. (Burlingame, CA).
Proteinase K and AmpliTaq GoldTM reverse transcriptase were
from Roche Molecular Biochemicals GmbH (Mannheim, Germany).
5,5,6,6'-Tetrachloro1,1',3,3'-tetraethylbenzimidazolecarbocyanide iodide (JC-1), MitoTracker®, Red AM, and ATP determination
kit were obtained from Molecular Probes (Eugene, OR). TRIzol reagent
was from Invitrogen. All other chemicals were of the highest
reagent grade and were purchased from Sigma.
Animals and Treatments--
Adult male Wistar rats (Rattus
rattus) were obtained from the Small Animal Facility of the
National Institute of Immunology (New Delhi, India). Wild type C57BL/6
(B6) and B6.MRL.lpr (B6-lpr,lpr) mice were obtained from the Jackson
Laboratory (Bar Harbor, ME) and maintained at the Small Animal Facility
of the National Institute of Immunology (New Delhi, India). Rats were
injected with different doses of DES (0.01, 0.1, and 1 mg of DES/kg of
body weight) every alternate day for varying periods of time with the
first day of injection being considered as day 0. After completion of
each treatment schedule, rats were killed by CO2
asphyxiation and testes were collected for various studies. Similarly,
adult 8-week-old lpr mice were treated with DES at 1 mg/kg body weight
for 7 days and were killed as above. For testosterone supplementation
studies, testosterone was administered at the dose of 30 µg/day for
appropriate time periods.
Preparation of Cells, Viability, and DNA
Analysis--
Spermatogenic cells were prepared according to Meistrich
et al. (18) with slight modifications as described
previously (19). Briefly, decapsulated rat testes were finely chopped
in spermatogenic cell medium (Ham's F-12/Dulbecco's modified Eagle's
medium, supplemented with 14.2 mM sodium bicarbonate, 10 mM sodium pyruvate, 2 mM
L-glutamine, 1 mM sodium pyruvate,
10
7 M testosterone, and 1% BSA) and the
cells were filtered through Nitex mesh (1000 and 20 µm), mira
cloth, and glass wool sequentially. After centrifugation, flow
cytometry was used to check the purity of the cellular preparation as
described earlier (19). FLOW JO software (Tree Star, Inc., Stanford,
CA) was used for DNA analysis.
Cell viability was checked by propidium iodide (PI) staining at a final
concentration of 5 µg/ml. PI-stained cell suspensions were analyzed
immediately on Coulter EPICS® ELITE ESP Flow Cytometer
(Coulter Corp., Miami, FL). DNA analysis of spermatogenic cells was
performed according to the procedure of Blanchard et al.
(20). Briefly, cells from treated and control animals were resuspended
in 50 mM PBS and chilled ethanol in a 1:1 ratio and
incubated on ice for 10 min. After washes, RNase treatment was given at
37 °C for 30 min and PI (50 µg/ml) was added to the cells, which
were analyzed in a flow cytometer as described above.
Preparation of Mitochondria and Cytosol--
Spermatogenic cell
mitochondria were prepared according to Graham and Higgins (21).
Briefly, 108 spermatogenic cells were suspended in
mitochondrial isolation buffer (150 mM sucrose, 10 mM succinate, 5 mM potassium phosphate, 10 mM HEPES-KOH, pH 7.4, 0.1% BSA) and lysed by nitrogen
cavitation (450 p.s.i. for 30 min at 4 °C with constant stirring).
The lysate was centrifuged sequentially at 500 and 2,500 × g for 10 min at 4 °C to remove unbroken cells and nuclei,
respectively. The resulting supernatant was centrifuged at 20,000 × g for 20 min at 4 °C to retrieve the pellet enriched
in mitochondria and stored in mitochondrial resuspension buffer (200 mM mannitol, 50 mM sucrose, 10 mM
succinate, 5 mM potassium phosphate, 10 mM
HEPES, pH 7.4, 0.1% BSA) for a few hours without significant loss of
the mitochondrial membrane potential (
m). The
supernatant obtained from the mitochondrial pellet was centrifuged at
100,000 × g for 1 h at 4 °C in an
OptimaTM XL-100K ultracentrifuge (Beckman, Palo Alto, CA)
to obtain the cell cytosol.
Tissue Processing, Terminal Deoxynucleotidyltransferase (TdT)
Enzyme-mediated dUTP Nick End Labeling (TUNEL) Assay, and
Immunostaining--
Testes from rats were fixed in Bouin's fixative
(saturated picric acid:formaldehyde:acetic acid in the ratio of 15:5:1)
for assessing morphological changes and in 4% formaldehyde (4%
formaldehyde in 50 mM PBS) for TUNEL assays. Tissues were
processed by standard procedures. Briefly, post-fixation wash and
dehydration in graded ethanol (50-100%) was followed by paraffin
embedding. Four-µm sections were cut with a Reichert Jung Microtome
1640 (Reichert, Germany) and transferred to albumin-coated slides. For
assessment of the morphological integrity of the seminiferous tubules,
tissue sections were stained with hematoxylin (0.5%) and eosin
(0.01%) after deparaffinization and hydration. Enumeration of
seminiferous epithelium stages I-IV, V-VI, VII-VIII, IX-XI, and
XII-XIV was carried out using a Nikon E600W upright microscope with a
40× objective. For each rat, at least 10 tubules per stage group were analyzed. These stages were identified according to the criteria proposed by Russell et al. (22) for paraffin sections. The
rate of spermatogenic cell apoptosis was expressed as the number of apoptotic spermatogenic cells per tubule.
Detection of DNA fragmentation by TUNEL staining was carried out as
described previously (19, 23) using a TUNEL assay kit according to
instructions from the manufacturer. Briefly, deparaffinized tissue
sections were treated with equilibration buffer (200 mM
potassium cacodylate, 25 mM Tris-HCl, pH 8.0, 0.2 mM DTT, 0.25 mg/ml BSA, 2.5 mM cobalt chloride)
for 10 min at room temperature followed by incubation with TdT buffer
containing nucleotide mix (50 µM dUTP-biotin, 100 µM dATP, 10 mM Tris-HCl, pH 8.0, 1 mM EDTA, pH 7.6) for 1 h at 37 °C. The
incorporation was visualized with avidin conjugated to horseradish
peroxidase (1:500) using a diaminobenzidine substrate kit, and sections
were counterstained with hematoxylin (0.5%) and visualized under a Nikon E600W microscope. At least 100 seminiferous tubules per rat from
four independent experiments were counted for TUNEL-positive nuclei.
For Fas-FasL immunostaining, fixed sections were deparaffinized and
blocked for endogenous peroxidase activity with 0.3%
H2O2 in PBS for 15 min and for nonspecific
binding with 10% normal goat serum for 30 min. Fas and FasL antibody
(1:100 dilution in 10% normal goat serum) incubation was carried out
at 4 °C, for 16 h. Appropriate blocking peptide at 4 µg/ml
concentration adsorbed with the antibody was added to the tissue
sections for identification of nonspecific staining. Anti-rabbit IgG
peroxidase (1:200) was used as secondary antibody for 1 h at room
temperature. After washing, color was developed with a diaminobenzidine
kit and sections were counterstained in hematoxylin (0.5%) for 30 s, dehydrated, and mounted in DPX mountant. For co-localization
of Fas-FasL, isolated spermatogenic cells fixed in 2% paraformaldehyde
were blocked for nonspecific sites with 10% normal goat serum
and permeabilized with 0.1% saponin prior to staining. The cells were
incubated with monoclonal antibodies against FasL (1:50) and Fas (1:50) tagged to phycoerythrin and fluorescein isothiocyanate (FITC), respectively. The dual staining was visualized with a confocal microscope (Zeiss LSM 510, Zeiss Inc., Thornwood, NY) by illuminating with a 488-nm argon ion laser at 15% power and images recorded through a band pass filter at 505-550 nm and a long pass filter at 560 nm.
For staining of Bax and cytochrome c, isolated spermatogenic
cells were fixed in 2% paraformaldehyde and stained for the respective antigens with anti-Bax and anti- cytochrome c antibodies
(1:50) for 1 h at room temperature. Secondary antibody tagged to
FITC was used at 1:200 dilution. For Bax-stained cells labeled with FITC, to identify the site of staining, cells were incubated with MitoTracker® Red AM, a mitochondria-specific dye, prior to
observation with a confocal system using same settings as above.
Stainings for MitoTracker® Red and Bax (green)
were overlapped and source of staining with anti-Bax antibody was
identified. Cytochrome c-FITC distribution in spermatogenic
cells was visualized at 540 nm using an E600W Nikon microscope.
Semiquantitative RT-PCR--
Total RNA was isolated from whole
testis and spermatogenic cells using TRIzol reagent. First strand
complementary DNA was made using 1-5 µg of total RNA in the presence
of AmpliTaq GoldTM reverse transcriptase and random primer.
After the RT reaction, 1 µl of incubation mixture was used as a
template for the subsequent PCR reaction. Primer sets used as a
template to obtain PCR products of FasL were
5'-AGCCCGTGAATTACCCATGTC-3' and 5'-TGCTGGGGTTGGCTATTTGCT-3' and for
-actin were 5'-AGGCATCCTGACCCTGAAGTAC-3' and
5'-TCTTCATGAGGTAGTCTGTCAG-3'. For semiquantitative analysis,
-actin
was used as an internal control, and was coamplified with FasL
messenger RNA by using
-actin primers and FasL primer (1 µl) in
the same mixture. All PCR reactions were performed for 30 cycles with
an annealing temperature of 55-65 °C in 1.5 mM
MgCl2.
Western Blot and DNA Analysis--
Electrophoresis on 12%
polyacrylamide gels, Western blots, and DNA extractions were carried
out as described previously (19, 23).
Measurement of Mitochondrial Membrane Potential
(
m) and ATP Levels--

m was
estimated using JC-1 as a probe according to the method of Dey and
Moraes (24) with slight modifications. JC-1 is a cationic mitochondrial
vital dye that is lipophilic and becomes concentrated in the
mitochondria in proportion to their 
m; more dye
accumulates in mitochondria with greater 
m and
ATP-generating capacity. The dye exists as a monomer at low
concentrations (emission, 530 nm; green fluorescence) but at higher
concentrations forms J aggregates (emission, 590 nm; red fluorescence).
JC-1 was chosen because of its reliability for analyzing

m in intact cells, whereas other probes capable of
binding mitochondria show a lower sensitivity or a noncoherent behavior
because of a high sensitivity to changes in plasma membrane potential
(25). Briefly, cells after different treatments were collected and
incubated for 10 min with 5 µM JC-1 at 37 °C, washed,
and resuspended in media, and 
m was measured at 590 nm for J-aggregates and at 530 nm for J-monomer. The ratio of 530/590
nm was considered as the relative 
m value.
ATP was measured by a bioluminescence assay (26) using an ATP
determination kit. The assay is based on the requirement of luciferase
for ATP in producing light (emission maximum ~ 560 nm at pH
7.8). Briefly, cells (106) after different treatments were
resuspended in reaction buffer containing 1 mM DTT, 0.5 mM luciferin, and 12.5 µg/ml luciferase and gently mixed,
following which readings were taken in a luminometer (Lumicount,
Packard, CT). ATP standard curves were run in all experiments with
different concentrations of ATP, and calculations were made against the
curve and cellular ATP levels were expressed as n
mol/106 cells.
Glutathione Levels--
Quantification of oxidized glutathione
(GSSG) and reduced glutathione (GSH) in the samples was done using the
fluorescent probe o-phthalaldehyde (23, 27).
Assay of Caspase Activity--
Cell lysates (200 µg of
protein) were incubated with caspase buffer (50 mM HEPES,
pH 7.4, 100 mM sodium chloride, 10% sucrose, 1 mM EDTA, 0.1% CHAPS, and 100 mM DTT)
containing 100 mM fluorogenic peptide substrate,
acetyl-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin (Ac-DEVD-AFC)
at 33 °C. Apopain from caspase-3 assay kit was used as a positive
control. 7-Amino-4-trifluoromethyl coumarin release was measured with a
PerkinElmer Life Sciences LS-50B luminescence spectrometer at
excitation wavelength of 400 nm and emission wavelength of 550 nm.
Testosterone Assay--
Plasma levels of testosterone were
measured by radioimmunoassay according to Sufi et al. (28).
The sensitivity of radioimmunoassay was 2.7 pg/assay tube. The
intra-assay and interassay variations were 10 and 7%, respectively.
Protein Estimation--
Bicinchoninic acid assay was performed
in microtiter plates as described previously (23).
Statistical Analysis--
An unpaired two-tailed Student's
t test using T-EASE software (version 2.0; Institute for
Scientific Information, Philadelphia, PA) and analysis of variance were
used for statistical analyses. Data are reported as mean ± S.E.
from at least three separate experimental groups unless specifically
mentioned. Each experimental group consisted of 6 rats. Data sets were
said to be significantly different for p < 0.05 (*),
p < 0.01 (**), and p < 0.001 (***).
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RESULTS |
DES Treatment Results in the Loss of Spermatogenic Cells and
Decrease in Serum Testosterone Levels--
DES, a potent estrogenic
analogue, is known to disrupt spermatogenesis by the suppression of
pituitary gonadotropins, which leads to the inhibition of testosterone
production (8) leading to insufficient circulating and intratesticular
concentration of the hormone. This results in abnormal spermatogenesis,
as testosterone is vital for spermatogenic cell survival (4). Even
though the importance of estrogen-induced deregulation of
spermatogenesis in male infertility has been acknowledged, the
mechanism of DES-induced apoptosis in the adult testis has not been
worked upon. Therefore, we first examined the kinetics of DES-induced
cell death in the adult testis by using two different doses
administered for varying periods of time. We observed a loss of
testicular weight in treated animals in a dose- and
time-dependent manner (Fig.
1A), indicating a decrease in
the number of testicular cells. This observation corroborates previous
studies where DES treatment in rat (8, 29) and hamsters (10) have been
shown to reduce testis weight that was accompanied by cell death.
Because DES is known to interfere with testosterone levels (30), we
checked serum testosterone and found a significant decrease in
circulating testosterone in treated animals in comparison to controls,
although a total shut-off was not achieved (Fig. 1B). In
addition to the above two doses of 0.1 and 1 mg of DES, we used 0.01 mg
of DES to look at similar parameters, but no appreciable change in
either testicular weight or testosterone levels were found (data not
shown).

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Fig. 1.
Effect of DES on testis weight and serum
testosterone levels. A, bar graph
showing changes in testicular weight at different time points after
treatment with increasing doses of DES. Note that for the 1-mg DES
dose, day 22 is represented as opposed to day 31 for the 0.1-mg DES
dose. Testis size was too reduced to weigh by day 31 in the 1-mg DES
group (n = 12, from two different experiments).
B, serum testosterone levels were reduced by 50% from day 7 onward with both 0.1- and 1-mg doses of DES. (n = 6).
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Histological examination of sections from the treated and control
testis showed that spermatogenic cell loss was more severe with 1-mg
DES treatment as compared with the 0.1-mg DES-treated group (Fig.
2A, a-g), even
though reduction in testosterone achieved with both the doses was
similar. The loss in testicular weight and spermatogenic cells from the
testis corresponded to a time- and dose-dependent decrease
in mean cross-sectional area of seminiferous tubules (Fig.
2B, a and b). The seminiferous tubular
atrophy observed during exposure to DES was accompanied by important
modifications in seminiferous epithelium composition and morphology.
There was an apparent decrease in total number of spermatogenic cells,
with both the number of spermatozoa and spermatid having decreased. The
type of spermatogenic cells affected by DES-induced testosterone suppression is different from those affected by gonadotropin-releasing hormone antagonist-induced suppression of testosterone (31). In the
regimen of hormone administration that we used, the group treated with
1 mg of DES exhibited a severe depletion of spermatogenic cells by day
22, whereas treatment with 0.1 mg of DES showed a nearly comparable
depletion of spermatogenic cells only by day 31, showing that depletion
of spermatogenic cells was dependent on dose as well as time of
treatment. Testosterone treatment along with DES was able to restore
serum testosterone levels to control levels from day 7 onward (Table
I). However, as shown in Table I, the
cell viability was not restored to control levels until day 14. This
was possibly a result of the combined suppressive effect of
testosterone and DES on the pituitary at the onset of treatment
schedule. As testosterone levels in the serum and the testis increased
with subsequent administrations, repopulation of the seminiferous
epithelium took place. The approximate time for spermatids to reappear
in the seminiferous epithelium would be 12 days according to the rat
seminiferous epithelial cycle length (22); therefore, the restoration
of cell viability in the testosterone supplemented groups to control
values after 14 days was logical. Taken together, the above data show
the ability of DES to reduce serum testosterone levels and induce a
loss of testicular spermatogenic cells in a dose- and
time-dependent manner that is paralleled by a loss in
testicular weight. Because spermatogenic cell loss was more extensive
with 1 mg of DES as compared with 0.1 mg of DES, the 1-mg dose regimen
was chosen for further study on the mode of DES action on spermatogenic
cell death.

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Fig. 2.
Effect of DES on testicular cell types and
seminiferous tubule diameter. A, sections from
testis of rats treated with DES at different time points showing
changes in seminiferous tubular cell populations. a,
control; b-d, sections from testis collected from 0.1-mg
DES-treated rats on days 7, 14, and 31 after treatment. Note the loss
of spermatogenic cells in some tubules, whereas others show intact
seminiferous epithelium. e-g, sections from testes
collected from 1-mg DES-treated rats on days 7, 14, and 22. Note the
severe depletion of spermatogenic cells from the seminiferous tubules
in g. Bar represents 100 µm. B,
seminiferous tubular diameter determined from testis sections of rats
treated with 0.1 mg of DES (a) and 1 mg of DES
(b). n = 6.
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Table I
Testicular weight, spermatogenic cell viability, and testosterone (T)
levels in testosterone supplementation studies
n = 6. *, p < 0.05.
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DES-induced Spermatogenic Cell Death Is Apoptotic in
Nature--
Based on the above data, we carried out a flow cytometric
analysis of spermatogenic cells prepared from testis of treated and
control animals to confirm histological observations. A significant cell loss was observed by day 7 in the 1-mg group as compared with the
controls (Fig. 3A,
a), thus confirming cell loss observed in earlier
experiments. Because it is known that testosterone withdrawal results
in spermatogenic cell apoptosis (30) and in our model DES was
causing a reduction in testosterone levels as well as death of
spermatogenic cells, we next investigated whether DES-induced cell
death expressed apoptotic or necrotic phenotype. By day 7 of treatment,
there was a substantial increase in cells with sub-haploid DNA content
as compared with controls (Fig. 3A, b) with a
concomitant reduction of haploid cell population, but there was no
change in the diploid population, indicating that the haploid spermatid
population was primarily affected. To ensure that spermatogenic cell
death was indeed apoptotic, we examined other apoptotic phenotypes
like DNA fragmentation by labeling the fragmented DNA ends with
fluorescent labeled nucleotides (TUNEL assay) (19, 23). TUNEL assay
showed a significant number of spermatogenic cells staining positive in
the DES-treated groups (Fig. 3B, b-d) as
compared with controls (Fig. 3B, a). The number of apoptotic cells per seminiferous tubule on days 7 and 14 after injection were 1.13 ± 0.1 and 6.66 ± 0.8, respectively, as
compared with control values of 0.03 ± 0.001, indicating
increased apoptosis in the seminiferous tubules (Fig. 3C).
This average number, however, is not a true representation of a
particular tubule at a given stage of the seminiferous epithelial
cycle. Stage VII showed the maximal number of TdT-positive cells
(14 ± 2) (n = 25), the numbers in other tubules
being lower on day 7 (1.0 ± 0.01) (n = 25).
Biochemically, internucleosomal DNA strand breaks, which are detected
as a ladder pattern on agarose gel electrophoresis of DNA are regarded
as a hallmark of apoptosis (32). Genomic DNA prepared from
spermatogenic cells of animals after day 3 of DES treatment showed
distinct DNA laddering regarded as a hallmark of apoptosis (Fig.
3D, b) as compared with controls (Fig.
3D, a). Therefore, the above data showing
nucleosomal DNA laddering, increase in TUNEL-positive cells and changes
in the ploidy of spermatogenic cells clearly established the apoptotic
nature of spermatogenic cell death, particularly in the haploid cell
type as induced by DES.

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Fig. 3.
Induction of cellular apoptosis by DES (1 mg). A, a, viability of spermatogenic cells
in control and 1-mg DES-treated groups as determined by flow cytometry
using PI staining of cells as a parameter for cell death
(n = 12 from two different experiments); b,
bar graph showing relative cell population with
different DNA contents in total spermatogenic cell preparations, as
determined by flow cytometry (n = 12 from two different
experiments). Note that on day 7, there is a significant increase in
the cell population with subhaploid DNA content indicating spermatid
fragmentation. B, TUNEL labeling of sections from treated
and control rats. a, control section showing very few cells
positive for TUNEL (brown deposits);
b-d, sections from testis of 1-mg DES-treated rats showing
increase in TdT-labeled cells/tubule on days 7, 14, and 22. Bar represents 10 µm. C, number of TdT-positive
cells/tubule on different days of treatment as indicated in the figure.
Cells on day 22 of treatment with 1 mg of DES could not be counted, as
very few cells were present in the tubules. (Each experimental group
consisted of 6 rats, and at least 100 tubules per rat testis were
counted.) D, genomic DNA prepared from control
(a) and treated (b) rats showing the presence of
DNA ladder from day 3 of treatment. All results are representative of
three different experiments with 6 rats each.
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Increased Expression of FasL and Fas Occurs in Spermatogenic Cells
Undergoing Apoptosis in DES-treated Rats--
Because it was evident
that DES-induced cell death occurred via apoptosis, the expression of
possible apoptosis-inducing proteins was studied to dissect the
apoptotic pathway. Given the existing evidence of the involvement of
the Fas-FasL system in the testis, we first looked at the expression of
Fas and FasL after DES treatment. Fas, a 45-kDa member of the tumor
necrosis factor receptor superfamily (33), binds to its cognate ligand,
FasL, and initiates the apoptotic pathway (34). It is generally
accepted that, in the testis, FasL is expressed in Sertoli cells, which
precipitates cell death by engaging its cognate receptors on
spermatogenic cells (4, 9). One recent report suggests that FasL
mRNA is expressed in the spermatogenic cells only (16). Our studies
revealed a clear up-regulation of FasL in spermatogenic cells within
24 h of DES treatment as determined by Western blots and RT-PCR
with FasL specific antibodies and probes, respectively (Fig.
4A, a and
b). Similar up-regulation of Fas was also detectable by
Western blots of purified spermatogenic cell extracts (Fig.
4A, c). Interestingly, in isolated spermatogenic
cells from DES-treated animals on day 1, FasL and Fas showed
differential localization. Although Fas was distributed evenly
throughout the cell, FasL staining was associated with vesicular bodies
(Fig. 4B, a-f). It has been reported that
FasL is stored in special secretory vesicles in certain cell types
(35); therefore, it is possible that, during the early part of DES
exposure, FasL is localized to secretory vesicles in spermatogenic
cells. To further clarify the cell types expressing these proteins,
immunohistochemical studies were carried out with testis sections.
Clearly, considerable up-regulation of FasL was visible in
spermatogenic cells. A lower intensity staining was observed in
spermatocytes and round spermatids, but more intense staining was
detectable in the elongated spermatid cytoplasm by day 7 after
treatment (Fig. 5). Interestingly, the
cytoplasm of the elongated spermatids of stage V and stage VII of the
seminiferous epithelial cycle were most reactive, with residual bodies
in stage VII that are remnants of spermatid cytoplasm staining
intensely as well (Fig. 5, A (c-f) and
C (a and b)). This differential
staining for FasL in different stages of the seminiferous epithelial
cycle is possibly related to variations in cell population and
physiological status of different stages of the cycle. It is known that
expressions of proteins and hormone binding capabilities vary with the
stages of the seminiferous epithelium (36). It is possible that
spermatogenic cells of stage V and VII were more susceptible to
estrogen-induced changes in terms of FasL induction. Fas staining was
visible in all stages of the seminiferous epithelial cycle, being
particularly prominent in stages VII and XII in all groups of
spermatogenic cells (Fig. 5, B (c-f) and
C (c)). When sections of testis from animals
treated with DES along with testosterone were examined for Fas and FasL
immunostaining, it was observed that there was a distinct
down-regulation of staining of both Fas and FasL in the
testosterone-treated groups by day 14 (Fig.
6) as compared with DES only (Fig. 5). In
summary, the above data show that DES treatment selectively increases
FasL and Fas expression in spermatogenic cells, indicating an apoptosis
modulating capacity of the spermatogenic cells independent of Sertoli
cell control, and provides a model for the study of Fas-FasL
interaction in the most complex epithelium of the body. In addition,
testosterone supplementation clearly down-regulates the Fas-FasL
up-regulation induced by DES, showing that testosterone is important
for regulation of apoptotic signals to germ cells. The fact that
Fas is an absolute necessity in case of DES-induced apoptosis was
proven by studies with lpr mice, where DES was unable to induce changes
in lpr mice ((testicular weight: lpr mice control, 0.17 ± 0.003 g; lpr mice + 1 mg of DES, 0.18 ± 0.004 g), (no. of TdT-positive
cells/100 spermatogenic cells: lpr mice control, 1 ± 0.008; lpr
mice + 1 mg of DES, 0.8 ± 0.005)).

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Fig. 4.
Changes in Fas-FasL in response to DES
treatment. A, a, up-regulation of FasL
protein after 1 and 7 days of DES treatment as shown by Western blots
of spermatogenic cell extracts. b, total RNA was extracted
from whole testis and spermatogenic cells (germ cells) and analyzed by
RT-PCR using primers specific for FasL and actin. The figure shows
increase in FasL messenger RNA in whole testis and spermatogenic cells
of DES-treated rat testis as evident from RT-PCR using specific
oligoprobes. -Actin was used as an internal control. c,
increase in Fas protein in response to DES treatment on 1 and 7 days.
B, co-localization of Fas and FasL by immunostaining of
isolated spermatogenic cells from vehicle-treated rats
(a-c) and from DES-treated rats (d-f) as
determined with a confocal microscope. a and d,
visualization of red (FasL) staining; b and
e, visualization of green (Fas) staining;
c and f, visualization of overlap of
red and green staining. Note a different staining
pattern with Fas antibody (green) and FasL antibody
(red) showing that, although Fas is distributed throughout
the cell, FasL on day 1 of treatment is present within secretory
granule-like structures.
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Fig. 5.
Immunohistochemical staining of Fas-FasL in
testis sections of DES-treated and untreated rats. A,
staining with anti-FasL antibody. a, control at stage V of
the seminiferous epithelial cycle; b, control at stage VII
of the seminiferous epithelial cycle; c, day 1 of treatment
at stage V of the seminiferous epithelial cycle; d, day 1 of
treatment at stage VII of the seminiferous epithelial cycle;
e, day 7 of treatment at stage V of the seminiferous
epithelial cycle; f, day 7 of treatment at stage VII of the
seminiferous epithelial cycle; g, section stained with
anti-FasL antibody adsorbed with FasL peptide. B, staining
with anti-Fas antibody. a, control at stage V of the
seminiferous epithelial cycle; b, control at stage VII of
the seminiferous epithelial cycle; c, day 1 of treatment at
stage V of the seminiferous epithelial cycle; d, day 1 of
treatment at stage VII of the seminiferous epithelial cycle;
e, day 7 of treatment at stage V of the seminiferous
epithelial cycle; f, day 7 of treatment at stage VII of the
seminiferous epithelial cycle; g, section stained with
anti-Fas antibody adsorbed with Fas peptide. Bar represents
10 µm. C, a, close-up of stage V tubule showing
spermatids inside a Sertoli cell. Note Sertoli cell cytoplasm does not
stain strongly for FasL, but the cytoplasm of the elongated spermatids
do. b, close-up of a section through a stage VII tubule
showing intense FasL staining associated with the cytoplasm of
elongated spermatids and residual bodies. c, magnified image
of a section through stage XII tubule showing the cytoplasm of the
elongated spermatids staining intensely for Fas.
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Fig. 6.
Immunohistochemical staining of Fas-FasL in
testis sections of rats treated with DES and testosterone.
A, staining with anti-Fas antibody. a, control
(C); b, DES + testosterone at day 1;
c, DES + testosterone at day 7; d, DES + testosterone at day 14. B, staining with anti-FasL antibody.
a, control; b, DES + testosterone at day 1;
b, DES + testosterone at day 7; d, DES + testosterone at day 14. Note that there is a significant reduction in
Fas and FasL staining in testosterone supplementation groups as
compared with DES treatment only at day 14 (shown in Fig. 5).
Bar represents 10 µm.
|
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Fas-FasL Up-regulation Is Followed by Increased Caspase-8 Breakdown
and Activation of Caspase-9 and -3--
Taking a lead from the above
data, we sought to determine the downstream effectors of the Fas-FasL
pathway including aspartate-specific cysteinyl protease (caspases)
family (37, 38). Our studies show that caspase-8 is cleaved to its
active form by day 1 after treatment (Fig.
7A). At this point, there were
two possible pathways that the spermatogenic cells could enter into;
one was to activate caspase-3 directly through the active caspase-8,
and the other was to activate caspase-9 by involving the members of the
Bcl-2 family of proteins. In our studies, processing of caspase-9 known to activate caspase-3 (39) to its active form of 37 kDa (37) was
detected prominently from 8 h onward (Fig. 7B).
Caspase-3 cleavage was detectable on Western blots of treated
spermatogenic cell extracts from day 1 onward (Fig. 7C). To
further confirm the activation of caspase-3, we directly determined
caspase-3 activity in DES-treated cell extracts by monitoring the
release of 7-amino-4-trifluoromethyl coumarin from Ac-DEVD-AFC, a
substrate of caspase-3. Caspase-3 activity was higher on day 7 as
compared with day 1 (Fig. 7D). Having established that
caspase-3 is active after DES treatment, we next checked for the
cleavage of poly(ADP-ribose) polymerase (PARP), a known endogenous
substrate for caspase-3. Cleavage into an 85-kDa C-terminal breakdown
product was detectable on Western blots, confirming that caspase-3 was
acting on cellular substrates (40) (Fig. 7E). Existing
studies show that Fas expression on the cell surface of some cell types
is p53-dependent (41). Because p53 can be activated by
changes in DNA and estrogen is known to induce changes in DNA (42), p53
expression was checked, but no change was detectable (Fig.
7F). Therefore, the above studies showing a temporal
relationship between the activation of caspase-8, translocation of Bax
to mitochondria, release of cytochrome c, and subsequent
activation of caspase-3 provide a clear picture that exposure to potent
estrogens could induce spermatogenic cell apoptosis in a
caspase-dependent manner.

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Fig. 7.
Time-course analysis of activation of
caspases, proteolytic cleavage of PARP, and expression of p53 after
treatment with DES. A, cleavage of procaspase-8 on days
1 and 2 of treatment. B, cleavage of procaspase-9 at
indicated time points after treatment. C, cleavage of
procaspase-3 to its active form in spermatogenic cells after DES
exposure at indicated time points. D, increase in caspase-3
activity as shown by caspase activity assay using Ac-DEVD-AFC as
substrate with spermatogenic cell lysates prepared after 1 and 7 days
of DES treatment. E, cleavage of PARP protein to 85 kDa as a
measure of caspase 3 activity on day 1 and 2 after treatment.
F, no change in p53 expression in spermatogenic cells was
observed after DES exposure on days 1 and 2. Results are representative
of 6 animals.
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Induction of Apoptosis by Caspase-8 Is Amplified through the
Mitochondrial Release of Cytochrome c Preceded by Bax Translocation to
the Mitochondria--
Because Bax, a multidomain, proapoptotic member
of the Bcl-2 family, has been shown to be involved in the first wave of
spermatogenesis (17), we first checked for changes in Bax localization,
as pro-apoptotic members of the Bcl-2 family translocate from the
cytosol to the mitochondria resulting in the release of cytochrome
c (43). Fig. 8 (A
and B) shows the migration of Bax from cytosol to the mitochondria of spermatogenic cells after DES exposure, where there was
a gradual loss of the monomeric form of Bax from the cytosol and
accumulation of oligomeric Bax in the mitochondria. Cytochrome
c was released by 8 h into the cytosol from the
mitochondria (Fig. 9, A and
B). The involvement of mitochondria in the activation of
caspase-3 clearly shows that adult spermatogenic cells follow the
apoptotic pathway taken by type II cells in response to estrogen treatment.

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Fig. 8.
Bax translocation from cytosol to
mitochondria after DES exposure. A, a,
accumulation of Bax protein (oligomer) in mitochondria as shown by
Western blot with anti-Bax antibody. Note the increase in Bax signal,
8 h and 1 day after DES exposure. b, relocation of Bax
from cytosol after DES exposure as detected by staining with anti-Bax
antibody. Note the decrease in staining intensity with increasing time
of exposure to DES in the cytoplasmic preparation. B,
co-localization of Bax protein (green) and mitochondria
(red) in control cells (a-c) and treated cells
(d-f); a and d, detection of
red fluorescence (mitochondria); b and
e, detection of green fluorescence (Bax
staining); c and f, overlap of green
and red fluorescence showing yellowish
orange staining indicating areas of co-localization. Note
strong co-localization in treated cells (f) as compared with
control cells (c).
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Fig. 9.
Effect of DES on cytochrome c
release. A, a, translocation of
cytochrome c from the mitochondria to the cytosol as
analyzed by Western blot with anti-cytochrome c antibodies,
showing decreased intensity of staining from day 1 onward in
mitochondria; b, increase in cytochrome c content
from 8 h onward in spermatogenic cell cytosol. B,
staining of spermatids with anti-cytochrome c antibody. Note
that control spermatogenic cells show a punctate localization of
cytochrome c (green) near the periphery of cells,
whereas at 1 and 7 days after treatment, much of the distribution of
cytochrome c is diffused even though some punctate staining
is visible. Bar represents 5 µm.
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Early Changes Upstream of Cytochrome c Are Mitochondrial Potential
Loss and ATP Depletion--
Looking at changes upstream to cytochrome
c release, we checked for mitochondria related events like
changes in 
m and ATP generation. Alteration in the

m are known to be a major cause of precipitation of
apoptosis in many cell types (43). Measurement of mitochondrial
membrane potential was carried out using the fluorimetric dye, JC-1
which showed a drop in 
m within 4 h after the
first injection (Fig. 10A).
This drop in 
m did not change significantly until day
7 after treatment with DES (data not shown). Any disruption of
mitochondrial function would reflect in the levels of ATP generation,
which was confirmed by a gradual fall in the ATP levels that reduced
significantly by day 7 of treatment with DES (Fig. 10B).
These data are consistent with the fact that the apoptotic process
requires ATP and therefore a minimal level maintained until day 7 might
be sufficient for continuing cellular apoptosis (44, 45). Glutathione
being a major detoxification molecule within the cell and changes in
the level of GSH can contribute to 
m loss, we
investigated GSH and GSSG levels. However, in case of estrogen-induced
spermatogenic cell death, there was no change in GSH and GSSG levels
((GSH, n mol/106 cells: control, 0.97 ± 0.08; day 1, 1.09 ± 0.1; day 7, 1.07 ± 0.09), (GSSG,
n mol/106 cells: control, 3.68 ± 0.2, day
1, 3.60 ± 0.4; day 7, 3.92 ± 0.5)), showing that GSH, even
though important in other models of germ cell death (19, 23) does not
seem to play any role in estrogen-mediated cell death. The above data
suggest that the loss of 
m that occurs by 4 h is
accompanied by a loss of cytochrome c and an increase in Bax
protein concentration in the mitochondria.

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Fig. 10.
Status of mitochondrial membrane potential
and ATP levels after DES exposure. A,
bar graph shows a significant decrease in
mitochondrial membrane potential, 4 and 8 h after DES treatment.
B, ATP concentration decreased significantly by day 7 of treatment.
|
|
 |
DISCUSSION |
Compelling evidence has accumulated on the detrimental effects of
estrogen-like chemicals present in the environment on male reproductive
function. These chemicals include industrial pollutants like bisphenol
A and polychlorinated biphenols, and pesticides like DDT, methoxychlor,
or chlorodecone. The extent of exposure to these chemicals on members
of a population differs as occupations in agriculture, petrochemicals,
and the construction industry entail higher exposure. Because estrogen
receptors are present in the pituitary and spermatogenic cells (1-3),
estrogen-like chemicals can act as agonists or antagonists for the
hormone and interfere with spermatogenesis. The DES-induced
spermatogenic cell apoptosis model was ideal to study the mechanism of
estrogen-induced spermatogenic cell death, as DES can mimic estrogen
action and has also been widely used as a model estrogen to study the
effects on the neonatal male rat reproductive tract (8, 9).
The finding that primarily haploid spermatogenic cells were undergoing
apoptosis in response to DES is in contrast to what is found in other
toxin-induced cell death models, where the diploid population of
spermatocytes are most affected (13, 14). Interestingly, immunoneutralization of follicle-stimulating hormone (46) and treatment
with gonadotropin antagonist (47), which suppress serum testosterone
levels like DES, precipitate apoptosis primarily in spermatogonia,
spermatocytes, and, to a lesser extent, spermatids. Clearly, in case of
DES treatment, the spermatids are mainly affected, suggesting that
estrogen-induced cell death primarily involves apoptosis of the haploid
population. This result is not surprising considering the fact that two
regulators of the cell death pathway, FasL and Fas, were overexpressed
in spermatids after DES exposure. Interestingly, the largest number of
apoptotic spermatogenic cells were visible in stage VII of the
seminiferous epithelial cycle, a stage were both FasL and Fas
expression were most pronounced and not in stage V or XII, where FasL
and Fas expression alone were most prominent. Therefore, the failure to
express FasL in a stage of the cycle where Fas was expressed or
vice versa showed substantially reduced number of cells
undergoing apoptosis. In other models of testosterone withdrawal in the
male, it has been shown that stage VII of the seminiferous epithelial
cycle was most susceptible in terms of apoptosis (48). Our study
clearly shows that the increased susceptibility of stage VII to cell
death is possibly a contribution of Fas-FasL interaction. The belief that Sertoli cells are the only source of FasL dominated thinking in
testicular biology (12); however, in the recent past it has been shown
that the spermatogenic cells have the ability to express both Fas and
FasL (16). The presence of Fas-FasL in testicular cells has been linked
to the ability of the testis to act as an immune privileged site and
maintenance of testicular homeostasis by Sertoli cells eliminating
spermatogenic cells. However, the recognition that both the Fas
receptor and its cognate ligand FasL can be expressed in spermatogenic
cells has led to the speculation that, apart from the above functions,
the expression of both these proteins on spermatogenic cells may
indicate an as yet unidentified function. Here, we demonstrate an
important novel function for the Fas-FasL system in the testis. We show
that estrogen exposure induces an increase in the expression of
spermatogenic cell Fas-FasL leading to the activation of components
downstream of Fas that indicate control of cell death within the same
lineage. More importantly, the observation that the number of apoptotic
cells is highest in seminiferous tubule stages where both Fas and FasL
expression was high reinforced the idea that these two proteins
expressed in spermatogenic cells are the main modulators of death.
Interestingly, in the female, strong FasL expression occurs in the
endometrium in estrogen-dependent late proliferative and
secretory phases (49). Additionally, in female rats, expression of FasL
protein and mRNA levels in ovarian cell increases upon estrogen
treatment (50). Therefore, our study shows for the first time that, in males as well, estrogen can induce expression of Fas-FasL system in the
spermatogenic cells. Apart from Fas-FasL up-regulation in cases of
acute exposure to estrogens, the expression of the receptor and ligand
may be important in the context of role of estrogens in normal
testicular functioning, as ERs have been identified on spermatogenic
cells but their function has not been identified. The absolute
requirement of Fas-FasL system in the event of DES-induced testicular
germ cell apoptosis comes from our studies with Fas-deficient lpr mice,
where we found that there was no reduction in testicular weight or
increase in TdT-positive cells after DES treatment in comparison to
controls. One other important finding that comes out of this study is
that spermatogenic cells are type II cells. The activation of caspase-8
is in consonance with the fact that engagement of Fas with FasL prompts
the formation of the death-inducing signaling complex composed by the
adaptor molecules Fas-associated death domain and procaspase-8,
followed by the release of active caspase-8. In many cell types (type
I), caspase-8 directly activates effector caspases, whereas in other
cell types (type II), Fas triggering induces insufficient disc
formation, inadequate to directly activate effector caspases yet
sufficient to initiate the mitochondrial apoptotic machinery (intrinsic
pathway) that in turn activates effector caspases (51). Our data
clearly show that the mitochondria is involved in DES-induced death, as
shown by the translocation of Bax from the cytosol to the mitochondria causing the release of cytochrome-c and loss of 
m.
This identification of spermatogenic cells as type II cells is
important, as therapeutic strategies that involve modulation of
Fas-mediated apoptosis must first identify the tissue to be determined
as type I or type II.
The intrinsic and extrinsic pathways are linked by the proapoptotic
members of the Bcl-2 family (43). In case of DES treatment, Bax protein
translocation to mitochondria occurred after caspase-8 was activated. A
role for Bax has been predicted in germ cell apoptosis using different
models, but predictions of involvement of Bax have largely been made
based on cellular localization (17, 52) or from Western blots of whole
testis extracts (53). Using purified spermatogenic cells, our study
clearly demonstrates that there is a translocation of Bax from the
cytosol to the mitochondria resulting in the release of cytochrome
c. In certain cell types, cytochrome c release is
accompanied by a fall in mitochondrial potential (54) and our studies
show that in spermatogenic cells the 
m fall is
preceded by a transient increase in reactive oxygen species
(data not shown) and followed by a fall in ATP levels.
Taken together, the data essentially illustrate the importance of the
Fas-FasL system in spermatogenic cell death in the event of estrogen
exposure. Furthermore, this study establishes germ cells as type II
cells that are able to utilize the extrinsic and intrinsic apoptotic
pathways, the link between the two pathways being the proapoptotic
protein Bax in the event of estrogen exposure. We also postulate that
normal testicular homeostasis may involve the Fas-FasL system to
maintain proper spermatogenic cell number, which may be under estrogen regulation.
 |
ACKNOWLEDGEMENTS |
We are grateful to G. S. Neelaram for
assistance in immunocytochemistry and to T. Nagarjuna for help in flow cytometry.
 |
FOOTNOTES |
*
This work was supported by grants to the National Institute
of Immunology from the Department of Biotechnology, Government of
India.The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
To whom correspondence should be addressed. Fax:
91-11-616-2125; E-mail: cshaha@nii.res.in.
Published, JBC Papers in Press, December 10, 2002, DOI 10.1074/jbc.M209319200
 |
ABBREVIATIONS |
The abbreviations used are:
ER, estrogen
receptor;
ED, endocrine disruptor;
DES, diethylstilbestrol;

m, mitochondrial membrane potential;
JC-1, 5,5,6,6'-tetrachloro1,1',3,3'-tetraethylbenzimidazolecarbocyanide
iodide;
PI, propidium iodide;
TdT, terminal
deoxynucleotidyltransferase;
TUNEL, terminal
deoxynucleotidyltransferase enzyme-mediated dUTP nick end labeling;
FITC, fluorescein isothiocyanate;
GSH, reduced glutathione;
GSSG, oxidized glutathione;
Ac-DEVD-AFC, acetyl-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin;
PARP, poly(ADP-ribose) polymerase;
DTT, dithiothreitol;
BSA, bovine
serum albumin;
PBS, phosphate-buffered saline;
RT, reverse
transcription;
CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid.
 |
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