Mol. Hum. Reprod. Advance Access originally published online on October 1, 2004
Molecular Human Reproduction 2004 10(11):825-834; doi:10.1093/molehr/gah099
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Somatic cell apoptosis markers and pathways in human ejaculated sperm: potential utility as indicators of sperm quality
1The Jones Institute for Reproductive Medicine, Department of Obstetrics and Gynecology, Eastern Virginia Medical School, Norfolk, Virginia and 2Center for Pediatric Research and Department of Physiological Sciences, Eastern Virginia Medical School and the Children's Hospital of the King's Daughters, Norfolk, Virginia, USA
3 To whom correspondence should be addressed at: Eastern Virginia Medical School, Center for Pediatric Research, 855 W. Brambleton Ave. Norfolk, VA 23510, USA. Email: sbeebe{at}chkd.com
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Abstract |
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In this study we extended earlier work to determine whether sperm respond to somatic cell apoptotic stimuli and whether apoptotic phenotypes are significant indicators of human sperm quality. We evaluated ejaculated sperm from fertile donors and subfertile patients following purification of fractions of high and low motility. In unstimulated conditions, caspase enzymatic activity was higher in motile fractions from subfertile patients than in donors, and was higher in low motility fractions from both groups. Staurosporine, but not a Fas ligand or H2O2, significantly increased caspase activity, but only in high motility fractions. Procaspase-3, -7 and -9 and low levels of active caspase-3, -7 and -9 were identified by immunoblot analysis. Apoptosis-inducing factor (AIF) was present in all samples but poly ADP-ribose polymerase-1 (PARP-1) was not detected. Phosphatidylserine translocation was significantly increased only with H2O2 treatment. In ejaculates of both subfertile and fertile men, we demonstrated the presence and activation of several proteins that are key constituents of apoptosis-related pathways in somatic cells, which may serve as markers for sperm quality.
Key words: caspase activity/apoptosis/apoptosis-inducing factor/infertility/phosphatidylserine translocation/sperm
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Introduction |
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Apoptosis is a well-characterized mechanism that allows eukaryotes to eliminate unneeded, senescent, or aberrant somatic cells (Kerr et al., 1972
). In general, somatic cell apoptosis can be induced through extrinsic mechanisms acting at the plasma membrane, mitochondrial or nuclear levels (Thornberry and Lazebnik, 1998; Gottlieb, 2001). The plasma membrane-dependent mechanism is typified by the interaction of the Fas receptor (CD95) and a Fas ligand that can proceed through two pathways characterized as type I and type II pathways (Scaffidi et al., 1998). The type I pathway is mitochondria independent, involving an adaptor protein to recruit caspase-8 to the cytoplasmic domain of the Fas receptor to form a death-induced signalling complex and activation of caspase-8. The type II pathway is mitochondria dependent, involving the truncation of Bid and the release of cytochrome c from the mitochondria, which binds to Apaf-I with ATP and caspase-9 to activate the latter by holoenzyme formation in the apoptosome. Activation of either initiator caspase-8 or -9 causes the activation of executioner caspases such as caspase-3, -6 and/or -7 (Thornberry and Lazebnik, 1998).
The identification of apoptosis in individual cells depends upon the presence of several different markers because a single reliable apoptosis phenotype may not be typical of all cell types. Perhaps one of the best markers for apoptosis is the presence of elevated caspase activity, but caspases may serve other functions unrelated to apoptosis (Martinez et al., 2002; Tesarik et al., 2002; Schwerk and Schulze-Osthoff, 2003). One well-characterized caspase-dependent marker of apoptosis is the cleavage and inactivation of poly-ADP-ribose polymerase (PARP), an enzyme involved in DNA repair. DNA fragmentation has been widely used as an apoptosis marker (Oberhammer et al., 1993; Nagata, 2000) caused by the activation of caspase-activated DNase (CAD). A caspase-independent factor released from the mitochondria, apoptosis-inducing factor or AIF, translocates to the nucleus where it also participates in DNA fragmentation (Susin et al., 1999; Daugas et al., 2000). Another cellular marker of apoptosis is phosphatidylserine translocation from the inner to the outer leaflet of the plasma membrane (Martin et al., 1995). During apoptosis, phosphatidylserine, normally present on the cytoplasmic face of the plasma membrane, is allowed to migrate to the outer leaflet. Here the exposed phosphatidylserine marks the cells for destruction by phagocytes (Fadok et al., 1992, 2001; Hoffmann et al., 2001), which occurs before loss of plasma membrane integrity to avoid inflammation, pain, and scarring.
At the nuclear level, the genome contains genes that are transcribed as a response to apoptotic stimuli. For example, p53 functions normally as a regulator of the cell cycle and a tumour suppressor in vivo. Following DNA damage, p53 induces apoptosis by up-regulation of the expression of the pro-apoptotic Bax gene and simultaneous down-regulation of Bcl-2 expression, a sensitive regulator–inhibitor of apoptosis (Selivanova and Wiman, 1995).
At the cytoplasmic level, several stimuli, including the activation of the mitochondrial membrane Bax, lead to release of cytochrome c (reviewed in Gottlieb, 2001). In the cytosol, cytochrome c stimulates a cascade of events leading to activation of caspase-3. Other endogenous or chemical apoptosis inducers such as ceramide, ATP depletion and staurosporine may act directly on mitochondria (Garland and Halestrap, 1997; Chandra et al., 2002; Siskind et al., 2002) (this is a membrane-independent, mitochondrial-dependent, apoptotic pathway) to permeabilize the outer mitochondrial membrane and to release AIF and/or cytochrome c. AIF directly translocates to the nucleus where it provokes large-scale DNA fragmentation and initial chromatin condensation.
While apoptosis in somatic cells and in (testicular) spermatocytes and spermatids in vivo is well established, the presence and significance of apoptosis in ejaculated human sperm is still unresolved (Oehninger et al., 2003). The Fas pathway has been implicated in apoptosis of spermatocytes in the testis (Lee et al., 1997; Pentikainen et al., 1999; Francavilla et al., 2000) and Fas receptors have been documented on human ejaculated sperm and correlated negatively with sperm concentration (Sakkas et al., 1999, 2002). Recently, we demonstrated the presence of immunoreactive inactive procaspase-3 and active caspase-3 (by immunoblotting and immunofluorescence), as well as caspase enzymatic activity in human ejaculated sperm (using a fluorometric assay; Weng et al., 2002). Others have confirmed these findings (Paasch et al., 2003; Wang et al., 2003ab). A single report of the presence of intact PARP in semen samples has been published (Blanc-Layrac et al., 2000). Furthermore, ejaculated sperm have been shown to present DNA fragmentation (Aravindan et al., 1997; Aitken et al., 1998; Evenson et al., 1999; Barroso et al., 2000; Duru et al., 2000; Duran et al., 2002; Marchetti et al., 2002; Sakkas et al., 2002; Weng et al., 2002). Likewise, phosphatidylserine (PS) translocation has been demonstrated in sperm upon different experimental conditions (Glander and Schaller, 1999; D'Cruz et al., 1999; Barroso et al., 2000; Gadella and Harrison, 2000; Schuffner et al., 2001, 2002; Paasch et al., 2003; Wang et al., 2003ab). Nevertheless, this phenomenon has also been related to positive functional changes in sperm such as capacitation (Gadella and Harrison, 2002; De Vries et al., 2003).
The objective of this study was to determine whether apoptotic phenotypes are significant indicators of human ejaculated sperm quality and whether such markers respond to somatic cell apoptotic stimuli.
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Materials and methods |
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Subjects and semen preparation
The Eastern Virginia Medical School Institutional Review Board (IRB) approved these studies. Subjects were recruited from two groups: fertile donors and men consulting for infertility (patients). Donor semen samples were obtained from fertile men participating in the artificial insemination donor sperm programme at the Jones Institute for Reproductive Medicine. All patients were consulting for primary infertility at the Jones Institute and had primary infertility of
1 year. All men had a normal physical examination and were non-smokers. Patients with antisperm antibodies (direct immunobead testing) and/or with round cell concentrations >1 x 106/ml were excluded from these studies. Patients and donors collected semen by masturbation into a sterile specimen container after 2–5 days of sexual abstinence. All semen samples had negative cultures for microorganisms. Viability was determined using eosin-Y. In order to be able to perform multiple assays in the same sample, only ejaculates having >40 x 106/ml motile sperm post-liquefaction were included.
Sperm were separated using discontinuous Percoll with low endotoxin levels (Sigma, USA) gradient separation. The high and low motility fractions were obtained from the 90 and 45% Percoll layers respectively. Contaminating leukocytes were removed using paramagnetic beads coated with CD45 antibodies (Dynabeads M-450 CD45 Pan leukocyte; Dynal, Norway) in accordanceb with the manufacturer's instructions as previously described (Weng et al., 2002). The absence of leukocytes was confirmed by methodical examination of peroxidase-stained slides. Samples containing >1000 residual polymorphnuclear leukocytes (PMN)/ml were subjected to an additional Dynabead separation step prior to use in assays.
Fractions were washed and resuspended in HTF–HEPES medium (Irvine Scientic, Santa Ana, CA) with 0.3% human serum albumin to a concentration of 40–50 x 106 sperm/ml. Immunodepleted cell suspensions were used in all experiments. The purified fractions containing sperm with high (90% layers) and low (45% layers) motility were then incubated for a 4 h period under conditions designed to stimulate an apoptotic response. All incubations took place at 37°C in humidified air. Each fraction was divided equally between four groups: 10 mmol/l staurosporine (S-5921; Sigma), 1 mg/ml anti-Fas antibodies (Fas ligand) (Clones CH-11 and IPO-4; Kamiya Biomedical Company, USA) and untreated (control condition). Each aliquot was evaluated for sperm concentration (manually) and motility parameters (using a computerized semen analyser, CASA) (Oehninger et al., 1990). Staurosporine and Fas ligand doses were selected as per their well-characterized effects on somatic cells. The selected dose of H2O2 has been previously shown to affect various sperm functions in vitro without reducing viability (Duru et al., 2000).
Caspase enzymatic activity
Sperm extracts were prepared using lysis buffer as previously described (Parvathenani et al., 1998; Weng et al., 2002). Briefly, sperm fractions (20–50 x 106 total spermatozoa) were prepared by sonicating sperm fractions in a lysis buffer on ice. Following sonication, suspensions were centrifuged to remove insoluble particulate matter. Aliquots of sperm lysates were incubated with the well-characterized fluorogenic substrate Ac-DEVD-afc (N-acetyl-aspartate-glutamate-valine-aspartate-AFC, 7-amino-4-trifluoromethyl coumarin) for 45 min at 37°C and fluorescent emissions (excitation 400 nm and emission 505 nm) were measured. Caspase activity (the assay measures activity of caspase-3, -6, -7, -8 and -10) was expressed as femtomole of substrate cleaved/min/106 cells. Human neutrophils treated with 1 mmol/l cycloheximide and 10 mmol/l staurosporine were used as positive controls.
Immunoblotting
Protein immunoblots were performed on dedicated aliquots of both high and low motility sperm fractions. Sperm fractions containing 40–50 x 106 cells were washed twice by centrifugation (10 minutes, 37°C, 300 xg.) and the pellet resuspended in phosphate-buffered saline. After the second centrifugation step, 5x electrophoresis sample buffer was added directly to the pelleted sperm. Reconstituted samples were then boiled for 10 min at 95°C. Extract proteins were separated by sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDS–PAGE) using a Bio-Rad Mini-Protean 3 Electrophoresis System (165-3301 Bio-Rad Laboratories, USA). The proteins were transferred to ImmobilonTM-P PVDF transfer membrane (Millipore Corporation, USA) using a Bio-Rad SD Semi-Dry Transfer Cell (Bio-Rad) according to the manufacturer's instructions. Membranes were blocked according to the instructions provided by the antibody supplier. ECL western blotting detection kits and Hyperfilm ECL films (Amersham Pharmacia Biotech, USA) were used to visualize and record protein bands according to the manufacturer's instructions and previously published reports (Qin et al., 2001). Human PMN or Jurkat cells were used as positive controls.
Primary antibodies for several apoptotic markers in somatic cells were used. The antibodies included polyclonal anti-caspase-3 (556425), anti-caspase-7 (551237), anti-caspase-9 (550437), and anti-poly-ADP-ribose polymerase (PARP) (65196E) from BD Pharmingen; AIF rabbit polyclonal antibody (NT) X1109P was obtained from Exalpha Biologicals Incorporated (USA).
Simultaneous detection of phosphatidylserine translocation and cell viability by flow cytometry
Recombinant FITC-conjugated Annexin-V (human) was obtained from Alexis Biochemicals, catalogue 209-250-T300 (Alexis Corporation, USA). Fraction concentrations were adjusted to a total 1 x 106 sperm and prepared according to the manufacturer's instructions for cell suspensions. Ethidium homodimer-1 (Molecular Probes, USA) was used as a counterstain for cell viability Vermes et al., 1995). Sperm suspensions were evaluated on a Becton-Dickinson FACalibur bench-top flow cytometer (Becton–Dickinson, USA) through excitation with a 15 mW 488 nm air-cooled argon-ion laser. For each suspension, 15 000 sperm were analysed for Annexin-V binding and/or ethidium homodimer uptake. Emission data were collected and analysed using CellQuest software (Becton–Dickinson).
Based on criteria for somatic cells upon flow cytometry, normal intact sperm were identified as annexin-V– and ethidium homodimer– appearing in the lower left quadrant; intact sperm with phosphatidylserine exposed on the outer membrane were identified as annexin-V+ and ethidium homodimer– appearing in the lower right quadrant; sperm with compromised membranes and viability were identified as annexin-V+ and ethidium homodimer+ appearing in the upper right quadrant, and as annexin-V– and ethidium homodimer+ appearing in the upper left quadrant.
Statistical analysis
Caspase activity results (patients versus donors, high versus low motility fractions) were compared using Generalized Estimating Equations analysis (Jung and Ahn, 2003; Mirkin et al., 2003). Flow cytometry data were analysed by rank transformation percentages in each of three groups (Annexin-V-bound live cells, unlabelled live cells, and necrotic cells) from dot plots of the data. The transformed data were then analysed using a single factor analysis of variance (ANOVA) with post-hoc multiple comparisons tests. Data are presented as mean±SEM. P<0.05 was considered significant.
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Results |
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The range of sperm parameters of samples used in these experiments were: (i) donors: concentration 53–267 x 106/ml, progressive motility 58–89%, and normal morphology 7–16%; and (ii) patients: concentration 30–178 x 106/ml, progressive motility 9–79%, and normal morphology 2–10%.
Variable levels of caspase activity were demonstrated in patients and controls and in their respective fractions of sperm with high and low motility
Figure 1 presents results of caspase activity (DEVD-afc assay) in fertile donors and in subfertile patients, in unstimulated conditions (control) and upon stimulation with each of the three apoptosis agonists (Fas ligand, staurosporine or H2O2); Figure 1A presents results of purified fractions of sperm with high motility (90% Percoll layers) and Figure 1B presents results of purified fractions of sperm with low motility (45% Percoll layers).
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In the high motility fractions (Figure 1A), unstimulated (basal) caspase activity was significantly higher in patients (n=25) than in donors (n=9) (P=0.02). Staurosporine treatment significantly increased caspase activity compared with control conditions in donors (n=9, P<0.001) and in patients (n=21, P=0.003). Staurosporine-stimulated caspase activity was significantly higher in patients than in donors (P=0.02). Fas ligand (n=8 donors and n=10 patients) did not result in significant changes in caspase activity. H2O2 treatment did not result in any significant change in caspase activity (n=8 donors and n=12 patients). In donors, staurosporine treatment was significantly higher than that of Fas ligand (P=0.01); in patients, staurosporine treatment was significantly higher than Fas ligand (P=0.003) and H2O2 (P=0.03).
In the low motility fractions (Figure 1B), unstimulated (basal) caspase activity was not significantly different between patients (n=18) and donors (n=9) (P=0.4). None of the three apoptosis agonists resulted in significant changes in caspase activity (donors: n=8, n=8 and n=9, for Fas ligand, H2O2 and staurosporine respectively; patients: n=11, n=4 and n=14 for the same agonists respectively).
In unstimulated (basal) conditions, caspase activity was consistently higher in the low motility compared to the high motility fractions. In donors this difference was nearly a 2-fold increase (P=0.06) whereas in patients it also followed a non-significant trend (P=0.6). Although we aimed to examine in parallel the high and low motility fractions from each sample (control and agonist-treated conditions), this was not always possible as the recovery rate following gradient separation was occasionally low in some instances. This explains the discrepancy in the sample sizes analysed. Fas ligand did not result in significant changes in sperm motility; staurosporine, on the other hand, resulted in significantly decreased motility but had no impact on viability (data not shown).
Various caspase isoforms were present in ejaculated sperm
To determine more specifically which pro- and active caspase enzymes were present and activated in the purified sperm fractions, immunoblot analyses were carried out using several different antibodies to pro- and active caspase-3, -7 and -9. Confirming our previous results (Weng et al., 2002), procaspase-3 (32 kDa) was identified in all high and low motility fractions of patients (n=6) and donors (n=9) tested. This polyclonal antibody also recognizes active caspase-3 (17 kDa active form). Low motility fractions of a minority of samples from both patients and donors showed active caspase-3, corroborating our previous report (Weng et al., 2002) (data not shown).
Figure 2 shows representative caspase-7 immunoblots of a donor (Figure 2A) and three patients (panel B). Like caspase-3, caspase-7 is an effector caspase. Procaspase-7 typically migrates to 
35 kDa during SDS–PAGE. An intermediate caspase-7 protein typically migrating at 32 kDa is formed upon cleavage of the 35 kDa proenzyme, presumably due to the cleavage and removal of the small pro-domain (Duan et al., 1996a; Lippke et al., 1996; Denault and Salveser, 2003). The active enzyme is composed of 20 and 11 kDa subunits.
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Figure 2A presents the results of a representative donor (out of three tested donors) as follows (from left to right): mol. wt marker, unstimulated human leukocytes (positive controls), low motility and high motility fractions. Unstimulated leukocytes showed both the procaspase (35 kDa) and intermediate procaspase-7 (32 kDa). The 35 kDa band was readily visible in all samples, suggesting that procaspase-7 is present in normal fertile donors regardless of the level of motility. However, the high motility fraction showed only a faint band when compared to the low motility fraction.
Figure 2B presents the results of the three studied patients as follows: unstimulated Jurkat cells and staurosporine-stimulated Jurkat cells [human leukocyte cell line used as positive controls, showing the procaspase (35 kDa), intermediate (32 kDa) and 20 kDa active fragment of caspase-7], and the low and high motility fractions of each patient A, B and C, consecutively. Although caspase-7 was absent in the high motility fractions, the low motility fractions of all three patients demonstrated the presence of procaspase (35 kDa) and intermediate (32 kDa) procaspase-7. The basic sperm parameters of these three infertile men were: patient A, concentration 46.2 x 106/ml, motility 53.8% and morphology 3.5%; patient B, concentration 72.3 x 106/ml, motility 71.4% and morphology 2.5%; and patient C, concentration 145.5 x 106/ml, motility 73.9% and morphology 6%.
Caspase-9 is an initiator caspase in somatic cells. It is activated by a mitochondria-mediated mechanism involving the release of cytochrome c and the formation of an apoptosome with Apaf-1 and caspase-9 (Li et al., 1997). The procaspase-9 protein migrates to 46–48 kDa during SDS–PAGE. Procaspase-9 is processed into a 37 kDa large subunit and a 10 kDa small subunit in cells undergoing apoptosis (Duan et al., 1996b). Results of representative immunoblots with the caspase-9 antibody are shown in Figure 3. Sperm fractions from one donor and the same three patients were used for this experiment. Unstimulated and staurosporine-treated Jurkat cells were used as positive controls. Procasapse-9 was present in both control conditions, and staurosporine stimulation resulted in the formation of active caspase-9 (37 kDa). Procaspase-9 was present in the high motility fraction of the donor. In the three patients, procaspase-9 was present to varying degrees in the high and low motility fractions. However, active caspase-9 was confined to a single patient (B) and to the sperm fraction with low motility.
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PARP, a caspase substrate in somatic cells, was absent in human ejaculated sperm
PARP-1 is a well-characterized as a substrate for caspase in somatic cells and often is used as a somatic cell apoptosis marker. PARP-1 typically migrates to 116 kDa during SDS–PAGE. In apoptotic cells, PARP-1 is processed to 85 and 25 kDa fragments (Tewari et al., 1995
). No intact PARP-1 or caspase-cleaved PARP-1 was detected in sperm of donors (n=9) or patients (n=8). Neither did human leukocytes in the presence or absence of staurosporine. Figure 4 illustrates a typical experiment using a PARP-1 antibody. As expected, untreated Jurkat cells displayed a band in the expected position of 116–120 kDa (Figure 4, lane 1). Jurkat cells were stimulated to undergo apoptosis by 4 h incubation with two different anti-Fas antibodies (CH-11 and IPO-4, both from Kaimya Biomedical Company, USA) and staurosporine. As expected, a band of 86 kDa was seen (Figure 4, lanes 2, 3 and 4). On the same blot, a representative fraction of high sperm motility from a fertile donor treated or not with staurosporine, and an extract from a staurosporine-treated PMN were evaluated for the presence of PARP-1. Human neutrophils did not exhibit PARP-1 in untreated or under staurosporine treated conditions.
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AIF was present in human ejaculated sperm
AIF, a phylogenetically old flavoprotein, is a 67 kDa protein. In non-apoptotic somatic cells, AIF is located in the mitochondrial intermembrane space. Upon apoptosis induction, AIF translocates to the nucleus where it binds to DNA and promotes caspase-independent chromatin condensation (Susin et al., 1999
; Daugas et al., 2000). To determine whether AIF was present in sperm, three patients and one donor were tested by immunoblot analysis with an AIF antibody. Figure 5 shows that AIF was detected in Jurkat cell controls as well as in the sperm extracts from the donor and the three subfertile patients.
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Analysis of phosphatidylserine translocation in sperm fractions from fertile donors treated with somatic cell apoptosis stimuli
Annexin-V–FITC is a well-characterized fluorescent apoptosis marker for the presence of phosphatidylserine on the outer plasma membrane of somatic cells (Vermes et al., 1995
). To determine if live sperm exhibited Annexin-V–FITC binding in response to somatic cell apoptosis stimuli, high and low motility sperm fractions from fertile donors were exposed to anti-Fas antibody (Fas ligand), staurosporine and H2O2 for 2 and 4 h, incubated with Annexin-V–FITC and ethidium homodimer-1, and analysed by flow cytometry. Figure 6 illustrates a typical flow cytometric analysis of a fraction with high sperm motility under control or treated conditions for 2 and 4 h. In this representative experiment, 71% of untreated sperm (control, 4 incubation) exhibited low fluorescence for annexin-V–FITC and ethidium homodimer appearing in the lower left quadrant, indicating that sperm were mainly intact (alive) and had low levels of phosphatidylserine exposed on their outer membrane. Nine per cent of the sperm in this sample appeared in the lower right quadrant exhibiting higher levels of Annexin-V–FITC fluorescence but low levels of ethidium homodimer. These represented intact sperm (alive) with phosphatidylserine on their outer membranes. Approximately 20% of the sperm were in the upper quadrants exhibiting high levels of ethidium homodimer fluorescence, indicating that their outer membranes were not intact.
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When sperm were treated with staurosporine for 2 or 4 h, no significant changes were observed in the flow cytometric profiles compared to control conditions. In contrast, sperm treated with H2O2 shifted fluorescence for both markers in a time-dependent manner. After 2 h of treatment, only 18% of the sperm were in the lower left quadrant and nearly 50% appeared in the lower right quadrant exhibiting annexin-V–FITC fluorescence in the absence of ethidium homodimer. In a somatic cell profile, these cells would be considered apoptotic. The number of sperm with compromised membranes increased from 20% to >30%. In a somatic cell profile these cells would be considered necrotic. After 4 h of H2O2 treatment, the percentage of sperm in the lower right quadrant decreased and the percentage of sperm in the upper quadrants increased proportionally. Thus, H2O2 resulted in an initial shift from the lower left to the lower right quadrant, indicating an increase in the percentage of intact sperm with phosphatidylserine on their outer membranes, followed by a secondary shift from the lower right quadrant to the upper right and left quadrants, indicating the occurrence of membrane disruption. This demonstrated a time-dependent appearance of phosphatidylserine on the outer membrane followed by a loss of membrane integrity. This is typical of somatic cell apoptosis followed by secondary in vitro necrosis as apoptosis proceeds to loss of membrane integrity.
Table I shows responses of the purified fractions of sperm with high and low motility from various fertile donors treated with staurosporine, Fas ligand and two different concentrations of H2O2 after 4 h of treatment. The results are limited to percentage of live sperm that remained intact (ethidium homodimer–) and exhibited phosphatidylserine on their outer membranes (Annexin-V+). The data indicated that the high dose of H2O2 (200 mmol/l) induced the appearance of phosphatidylserine on the outer membranes both in the high and low motility fractions (P=0.01 versus controls). The lower H2O2 dose (25 mmol/l) and the other somatic cell apoptosis stimuli (Fas ligand and staurosporine) were without effect.
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Discussion |
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As an extension of previous work (Weng et al., 2002
), in this study we analysed apoptosis proteins and pathways in leukocyte-free ejaculated sperm, eliminating contributions of somatic cell apoptosis markers. Ejaculated sperm exhibited several apoptosis-related proteins, expressed caspase catalytic activity that could be elevated by some somatic cell apoptosis stimuli, and displayed some somatic cell apoptosis markers that were differentially observed in fertile donors and in infertile patients. These somatic cell apoptosis-related proteins and markers could (i) serve apoptosis death functions, (ii) serve other functions, including roles in fertility or infertility, or (iii) serve no function if they were remnants of previous pre-ejaculation stage activities.
To define analogous roles for somatic cell apoptosis-related proteins in ejaculated sperm, several phenotypic and biochemical events should be correlated with loss of sperm structure and/or function or with clinical infertility. In this study, several correlates were observed. First, caspase catalytic activity was higher in low motility fractions (dyskinetic and dysmorphic sperm) compared to high motility sperm fractions. Second, caspase enzyme activity was higher in the ejaculates of infertility patients than in fertile donors. Third, in high motility fractions, staurosporine induced caspase activity in fertile donors, but not in infertility patients. This is most likely because caspase activity was already elevated in patient ejaculates and/or perhaps due to the fact that highly motile sperm have more intact machinery, making them more susceptible to the induction. The question remains if these somatic cell apoptosis markers indicate sperm apoptosis or if they serve other functions. Given known functions for caspases (Thornberry and Lazebnik, 1998), it seems more likely that the presence of elevated caspase enzyme activity and other caspase markers in dyskinetic and dysmorphic sperm have some function or have served their functions. The elevated levels of active caspases may be related to poor sperm quality and could be a possible indicator of infertility. Whether this is an indication of apoptosis or some other form of sperm dysfunction requires further study.
In a previous study (Weng et al., 2002), inactive and active caspase-3, an effector caspase, were unequivocally identified in the mid-piece of ejaculated sperm and the presence of caspase-3 was confirmed in these studies. To further define the presence and potential roles for somatic cell apoptosis markers in ejaculated sperm, we analysed other potential immunoreactive apoptotic proteins in ejaculated sperm including caspase-3, -7 and -9, PARP, and AIF.
Profiles of caspase-7 immunoreactivity, another effector caspase, in fertile donors and infertility patients were particularly interesting. Immunoreactive procaspase-7 and its intermediate fragment were present in the low, but not in high, motility fractions of infertility patients. Furthermore, only very low levels of procaspase-7 and none of the intermediate fragment were present in high and low motility fractions from a typical fertile donor. The intermediate fragment of caspase-7 arises by the cleavage and removal of the pro-domain of the enzyme (Denault and Salvesen, 2003). The pro-domain has been proposed for initiator caspases to prevent activity in the procaspase during and after protein folding (Riedl et al., 2001). Thus, the presence of the intermediate form of caspase-7 may represent caspases that are primed for activity. The presence of procaspase-7 and the intermediate fragment in untreated human neutrophils, which undergo apoptosis as a terminal function in vivo and in vitro (Parvathenani et al., 1998), support this concept. This suggests that high levels of procaspase-7 and/or its intermediate fragment should be considered for further investigation as a potential marker for low sperm quality and/or infertility.
Active forms of caspase-3, -7 and -9 were present and confined to the low motility fractions and typically to patients' samples, suggesting that they may be indicative of poor sperm quality. The presence of immunoreactive active caspases was weakly demonstrated in some but not all sperm fractions whereas caspase catalytic activity was more readily demonstrated, albeit at levels lower than those found in somatic cells. Determination of catalytic activity with the pan-caspase substrate DEVD-afc is a considerably more sensitive caspase marker (Parvathenani et al., 1998; Morris et al., 2002), suggesting that the low immunoreactive presence of caspases in sperm extracts may be due to the low sensitivity of immunoreactive caspase proteins. It is also possible that low caspase activities could be due to the presence of caspases not readily detected by either of these methods, to a low cytoplasmic volume, or to the presence of caspase activity in a relatively small subpopulation of sperm. Furthermore, while the proteolysed, active fragment of caspase-9, an initiator caspase, was identified in at least one patient sample, the activation of caspase-9 occurs by holoenzyme formation with Apaf-1 and does not necessarily require cleavage processing. Finally, it is possible that low active caspases could be due to other functions that require tight regulatory control.
While caspase activity is well characterized for positive feed-forward cell disassembly during apoptosis in somatic cells, it has recently become evident that caspases are involved in functions other than apoptotic cell demise, including muscle cell differentiation (Fernando et al., 2002), T-cell proliferation (Kennedy et al., 1999) and T-cell function (Alam et al., 1999). In these functions, caspase activity would need to be highly regulated at low levels and/or in subcellular microenvironments. Thus, the low levels of caspase activity observed in ejaculated sperm could be involved in mechanisms other than cell death (Blanco-Rodriguez and Martinez-Garcia, 1999; Tesarik et al., 2002). It is tempting to speculate that, in addition to possible roles in infertility and/or cell death, caspases in ejaculated sperm could serve remodelling functions during capacitation and/or acrosome reaction, aid sperm penetration of the oocyte during fertilization, and/or have post-fertilization functions. If these putative functions were initiated prematurely, fertility could be affected. Whether caspase activities were initiated in testicular sperm or occurred as post-ejaculatory events, the presence of elevated caspase enzyme activity or immunoreactivity could compromise sperm function and quality, as observed in lower quality ejaculated sperm, and be related to infertility, as observed in infertility patients.
As an additional marker for caspase activity, a caspase substrate was analysed. PARP cleavage, a well-characterized marker for caspase activity and apoptosis in somatic cells, was not present in ejaculated sperm and could not be used as a potential apoptosis marker.
Another somatic cell apoptosis marker was analysed in ejaculated sperm. AIF, which we show here to be present in ejaculated sperm, could be important in DNA degradation since, in somatic cells, it translocates from the mitochondria to the nucleus and promotes caspase-independent chromatin condensation (Susin et al., 1999; Daugas et al., 2000). The implication of AIF would also explain the observations on apoptosis-related phenomena in late spermatids and sperm that may lack caspase activity but are rich in mitochondria (Tesarik et al., 2002).
In addition to evaluating apoptosis markers in untreated sperm, we incubated sperm with known somatic cell apoptosis agonists including staurosporine, H2O2, and anti-Fas. We observed that (i) staurosporine induced caspase activity but not translocation of phosphatidylserine; (ii) H2O2 increased phosphatidylserine translocation and weakly activated caspases, although the latter effect was not statistically significant; and (iii) anti-Fas did not affect any of these markers. Thus, somatic cell apoptosis markers appear to be agonist dependent and only partially correlated with putative apoptosis markers.
Staurosporine is well characterized to disrupt mitochondria (Tafani et al., 2001) and may result in apoptosis by stimulation of different pathways, including AIF release, disruption of Bax–Bcl equilibrium, generation of reactive oxygen species (ROS), and/or release of cytochrome c and Smac/Diablo (reviewed in Gottlieb, 2001). The latter two products may activate cytosolic caspases (Wolf et al., 1999). Interestingly, caspases may be localized in the intermembrane mitochondrial space in somatic cells (Mancini et al., 1998; Susin et al., 1999). Our results demonstrated that staurosporine induced caspase activation particularly in high motility fractions. Nevertheless, further studies are needed to decipher downstream effects and their relation to cellular apoptotic changes.
ROS including H2O2 have been shown to induce apoptosis and activate caspases in somatic cells (Stridh et al., 1998; Chiaramonte et al., 2001). Treatment of ejaculated sperm with H2O2 resulted in increased phosphatidylserine translocation. ROS have been reported to affect the function of the aminophospholipid transferase, resulting in externalization of phosphatidylserine (Gottlieb, 2001). In previous experiments we have shown that the same dose of H2O2 induced DNA fragmentation as examined by TdT (terminal deoxynucleotidyl transferase)-mediated dUDP nick-end labelling (TUNEL) (Duru et al., 2000). In the present experiments, H2O2 had only a marginal effect on caspase activation. On the other hand, Paasch et al. (2003) showed that deterioration of the sperm plasma membrane (characterized by phosphatidylserine externalization) was associated with activated caspases. However, externalization of phosphatidylserine has been related to other sperm conditions such as capacitation (Gadella and Harrison, 2002; De Vries et al., 2003). The correlation of somatic cell apoptosis markers with sperm morphology and biochemistry requires further analysis; however, based on results presented here, it cannot be concluded that externalization of phosphatidylserine secondary to H2O2 was the result of an apoptosis phenomenon, either caspase dependent or independent.
Whereas the significance of Fas-mediated pathways during apoptosis in testicular tissue is well documented (Lee et al., 1997; Pentikainen et al., 1999; Francavilla et al., 2000), treatment of leukocyte-free, ejaculated sperm with anti-Fas receptor did not result in any change in apoptosis markers. We therefore concluded that although Fas binding sites have been described in ejaculated human sperm (Sakkas et al., 1999), the receptor might be either non-functional or non-operative after ejaculation. Further studies are needed to unequivocally demonstrate the presence and functional response of Fas receptors in human ejaculated sperm.
In some somatic cells in which partly or totally caspase-independent apoptosis was observed, cell death was induced by oxidative stress (Carmody and Cotter, 2000; Kim et al., 2000; Krishnamurthy et al., 2000). Similarly, oxidative stress may play a role in apoptosis of human germ cells in which the apoptotic process induced by withdrawal of survival factors (serum or hormones) is attenuated by lowering partial oxygen pressure (Erkkila et al., 1999). In a recent study using testicular biopsies, Tesarik et al. (2004) demonstrated that germ cells that are tightly associated with Sertoli cells undergo caspase-dependent apoptosis and show phosphatidylserine externalization. Other germ cells that lack a tight association with Sertoli cells show caspase-independent DNA fragmentation and do not externalize phosphatidylserine. While it is not clear if these mechanisms are present in ejaculated sperm, based on the identification of AIF in ejaculated sperm, it is possible that AIF could be a candidate for a caspase-independent, mitochondrial-activated process in human sperm.
Sakkas et al. (2002) investigated the relationship between nuclear DNA damage (assessed by TUNEL) and key apoptotic proteins, including Fas, Bcl-x and p53. In agreement with our results, apoptotic markers did not always exist in unison, but nonetheless, there was some correlation between the higher content of apoptosis markers and the presence of poor sperm parameters. The authors speculated that there might be a population of sperm that have escaped programmed cell death and express various apoptotic markers, or an ‘abortive apoptosis’ (Sakkas et al., 1999), which might be linked to defects in the cytoplasmic remodelling during late spermatogenesis. In agreement, we speculate that there might be a subpopulation of sperm that have initiated an apoptosis-like programme that may have bearing on fertility and infertility regardless of whether this is defined as sperm apoptosis.
Not all studies have observed apoptosis phenotypes in ejaculated sperm. A recent study could not demonstrate the appearance of phosphatidylserine externalization or DNA fragmentation during 4 and 24 h in vitro incubations of ejaculated sperm from fertile donors (Lachaud et al., 2004). After 24 h, sperm viability and motility declined significantly, suggesting sperm demise without exhibiting phosphatidylserine externalization or DNA fragmentation. However, this study did not evaluate sperm from infertility patients, use somatic cell apoptosis stimuli, or analyse caspase activity, which is indicated as a highly sensitive somatic cell apoptosis marker in the present study. Based on results shown here, caspase activity may be an important marker for sperm quality in at least a subset of sperm from fertile and/or infertile patients.
Taken together, our observations suggest that an apoptosis-like phenotype can be found in ejaculated sperm similar in some ways, but not all, to those observed in somatic cells. This occurs more frequently in infertile men with abnormal sperm parameters and also in the purified fractions of sperm obtained from the low gradient centrifugation layers. Examples of this conserved apoptotic phenotype include (i) elevated caspase activity especially in dyskinetic and dysmorphic sperm, (ii) externalization of phosphatidylserine in H2O2-induced, but not staurosporine-induced, sperm demise, and alterations of mitochondrial membrane potential (early markers) as well as DNA damage (late marker); and (iii) the identification of several of the well known key proteins involved in somatic cell apoptosis and their activation by defined agonists, which provides a mechanistic platform for the occurrence of pathways that may lead to sperm dysfunction, infertility, and/or death. While overall the data do not clearly indicate that ejaculated sperm undergo apoptosis, they do suggest that these markers might have clinical relevance as some of them could potentially be used as diagnostic tools to predict sperm dysfunction and male infertility.
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