Molecular Human Reproduction, Vol. 8, No. 11, 984-991,
November 2002
© 2002 European Society of Human Reproduction and Embryology
Testis and spermatogenesis |
Caspase activity and apoptotic markers in ejaculated human sperm
1 The Jones Institute for Reproductive Medicine, Department of Obstetrics and Gynecology and 2 The Center for Pediatric Research, Eastern Virginia Medical School, Norfolk, VA, USA and 3 Reproductive Endocrinology and Infertility Division, Department of Obstetrics and Gynecology, Mackay Memorial Hospital, Taipei, Taiwan, Republic of China
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Abstract |
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The objectives of this study were to determine if human ejaculated sperm exhibit active caspases and if caspase-dependent apoptosis markers are identifiable. Sperm from fertile donors and infertile patients were examined after gradient separation into leukocyte-free fractions of high and low motility. Sperm were evaluated for motion parameters, morphology, caspase activation, and apoptosis markers including phosphatidylserine (PS) translocation (annexin V binding) and DNA fragmentation (TUNEL). Active caspase-3 was detected by immunofluorescent microscopy in a small proportion of sperm in situ, in fractions of high and low motility sperm of patients and donors, but low motility fractions had significantly higher numbers of positive sperm. Immunoblot analysis detected inactive procaspase-3 (32 kDa) in all fractions of low sperm motility from patients and donors, while active caspase-3 (17 kDa) was only detected by immunoblotting in a limited number of low motility fractions from patients and in even fewer fractions from donors. Caspase enzymatic activity, as measured using the fluorogenic substrate DEVD-afc, was higher in patients than in donors in both low and high motility fractions. Annexin V staining and DNA fragmentation were detected in a proportion of sperm, with a higher frequency in the low motility fractions. A significant positive correlation between in-situ active caspase-3 in the sperm midpiece and DNA fragmentation was observed in the low motility fractions of patients, suggesting that caspase-dependent apoptotic mechanisms could originate in the cytoplasmic droplet or within mitochondria and function in the nucleus. These data suggest that in some ejaculated sperm populations, caspases are present and may function to increase PS translocation and DNA fragmentation.
apoptosis/caspase-3/DNA fragmentation/phosphatidylserine translocation/sperm
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Introduction |
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Apoptosis is a mechanism that enables metazoans to control cell number in tissues and to eliminate individual cells that threaten the animal's survival. Apoptosis also plays an essential role in the processes of gamete maturation and embryogenesis, contributing to the appropriate formation of various organs and structures. Studies in animal models have demonstrated that apoptosis is the underlying mechanism of germ cell death during normal spermatogenesis (Sinha-Hikim et al., 1998
). Relatively high rates of apoptosis have been observed in testicular biopsies from infertile men with varying degrees of testicular insufficiency, using in-situ end-labelling (TUNEL) and morphometric criteria (Jurisicova et al., 1999).
Recent reports have demonstrated that ejaculated sperm from infertile men show ultrastructural damage, an unusually high incidence of DNA fragmentation and plasma membrane translocation of phosphatidylserine (PS), all of which are typically considered to be signs of apoptosis in somatic cells (Gorczyca et al., 1993; Baccetti et al., 1996; Aitken et al., 1998; Lopes et al., 1998; Barroso et al., 2000). These effects can also be induced by defined in-vitro sperm treatments (D'Cruz et al., 1999). The alterations are typical of apoptosis in other cells and are suggestive of apoptosis in ejaculated sperm (Gavrieli et al., 1991; Kerr, 1993; Martin and Green, 1995; Vermes et al., 1995; Barroso et al., 2000). Nonetheless, these sperm may be considered normal using routine semen analysis. Under certain conditions (in vivo or after in-vitro therapy, particularly ICSI), these sperm could possibly carry a damaged genome into the oocyte, resulting in serious consequences (Bowen et al., 1998; Bonduelle et al., 1999).
It is now believed that a central component of apoptotic machinery involves one or more members of a family of aspartic acid-directed cysteine proteases called caspases (Thornberry and Lazebnik, 1998). In healthy cells, caspases are expressed as inactive proenzymes (~30 kDa) that contain three domains: an NH2-terminal domain, a large subunit (~20 kDa) and a small subunit (~10 kDa). These enzymes participate in a cascade triggered in response to pro-apoptotic signals and culminate in proteolysis of proteins essential for cell homeostasis, ultimately resulting in death of the cell. Caspase-3 is the main executor within this apoptotic cascade. Understanding the mechanism of caspase regulation is intimately linked to the ability to rationally manipulate apoptosis for therapeutic gain (Thornberry and Lazebnik, 1998; Brill et al., 1999).
The objectives of this study were: (i) to investigate the presence of active caspase-3 in ejaculated human sperm; (ii) to determine whether the appearance of typical caspase-dependent apoptosis markers could be identified in human ejaculated sperm; and (iii) to determine whether apoptosis markers can be used in therapeutic tools for infertility. Caspase activity was demonstrated by three distinctly different methods, including: (i) an in-vitro fluorometric assay for hydrolysis of Ac-DEVD-afc, which is a good substrate for several caspases including caspase-3, -6, -7 and -8 (Thornberry and Lazebnik, 1998); (ii) immunoblot analysis for detection of inactive (32 kDa) and active (17 kDa) caspase-3; and (iii) immunocytochemistry for identification of active caspase-3 in situ. Results of caspase activation were compared with the presence of a relatively early apoptosis marker, plasma membrane translocation of PS as monitored by annexin V binding, and a relatively late marker, DNA fragmentation as assessed by TUNEL, in purified fractions with high and low sperm motility from infertile patients and fertile donor controls.
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Subjects
The Institutional Review Board of the Eastern Virginia Medical School approved these studies. Semen was obtained from men (25–43 years old) consulting for infertility and who were undergoing basic semen analysis at our andrology laboratory. Patients had primary infertility of at least 1 year duration. All were non-smokers and had a normal physical examination. To ensure sufficient sperm to perform all planned measurements, patients with a total concentration <20x106/ml motile sperm in the liquefied sample were excluded from the study. In addition, fertile men participating in our artificial insemination programme (donors) were studied as controls. Patients and donors collected semen by masturbation into a sterile specimen container after 2–5 days of sexual abstinence. All semen samples contained <1x106 round cells/ml (peroxidase positive) and cultures for micro-organisms were negative.
Sperm concentration, motility and morphology
All semen samples were allowed to liquefy for 30 min at 37°C followed by assessment of sperm parameters. Sperm concentration and motion parameters were assessed using the HTM-IVOS semen analyser version GS 771 (Hamilton Thorne Research, Beverly, MA, USA) and manually monitored as appropriate. Sperm motion analysis was performed with fixed parameter settings as previously described (Oehninger et al., 1990). The following motion parameters were measured: percentage progressive motility, average path velocity, straight line velocity, curvilinear velocity, amplitude of lateral head displacement, linearity, beat frequency and rapid cells (percentage sperm with velocity >50 µm/s). Sperm morphology percentage normal forms and percentage cells with cytoplasmic droplets were assessed using strict criteria after staining the smear of semen with Diff-Quik (Dade AG, Dudinger, Switzerland).
Preparation of samples using discontinuous Percoll gradient separation
Sperm were separated using discontinuous Percoll (Sigma, St Louis, MO, USA) gradient separation (90 and 45% layers). Modified human tubal fluid (HTF)–HEPES (Irvine Scientific, Santa Ana, CA, USA) supplemented with 0.3% (w/v) human serum albumin (HSA; Irvine) was used as a dilution medium for Percoll. Upon completion of Percoll separation, purified populations of highly motile (from the 90% layer) and poorly motile (from the 45% layer) sperm were recovered. Contaminating leukocytes were removed using paramagnetic beads coated with CD45 antibodies (Dynabeads M-450 CD45 Pan leukocyte; Dynal, Oslo, Norway) in accordance with the manufacturer's instructions and previously published studies (Krausz et al., 1992; Aitken et al., 1996). The absence of leukocytes was confirmed by methodical examination of peroxidase-stained slides and measurement of reactive oxygen species in the presence of FMLP (N-formyl-met-leu-phe). Samples were washed and resuspended in HTF–HEPES medium with 0.3% HSA to a concentration of 40–50x106 sperm/ml.
Immunofluorescent detection of activated caspase-3 in sperm
To detect active caspase-3, we used a purified rabbit anti-active caspase-3 monoclonal antibody (PharMingen, San Diego, CA, USA), which specifically recognizes the active form of caspase-3 in human cells and does not recognize the proenzyme (Weil et al., 1998). Processed sperm samples were washed with phosphate-buffered saline containing sodium azide (0.5%) and phenylmethylsulphonyl fluoride (0.1 µmol/l) twice. Samples were prepared as previously described (Weil et al., 1998). Bound antibodies were visualized using a FITC-conjugated goat anti-rabbit immunoglobulin-specific polyclonal secondary antibody (PharMingen). Human neutrophils isolated from whole blood through a Percoll gradient were used as positive controls. Evaluation for active caspase-3 in sperm was completed in a blinded fashion by two evaluators at a magnification of x1000 using an epifluorescent microscope equipped with phase-contrast optics (Eclipse 600; Nikon, Melville, NY, USA) and using a digital camera with a high pressure mercury lamp power supply (SPOT RT, software version 3.2; Diagnostic Instruments, Augusta, GA, USA). At least 500 sperm were examined per slide, in duplicate slides. The intra- and inter-observer coefficients of variation were 6 and 4% respectively.
Immunoblot analysis for active caspase-3
Sperm (40–50x106) were centrifuged to a pellet, placed on dry ice (5–10 min) and lysed in Tris buffer with 0.5% Triton, pH 7.5, containing general protease inhibitors (Parvathenani et al., 1998). The sperm suspension was vortexed and sonicated before centrifugation to remove particulate matter. The supernatant was removed, mixed with 5x loading buffer and boiled. Samples were separated on a 10% sodium dodecyl sulphate/polyacrylamide gel and transferred to Immobilon-P PVDF transfer membranes (Millipore Corp., Bedford, MA, USA). Membranes were blocked with 5% non-fat dry milk and then incubated with a polyclonal rabbit anti-caspase-3 antibody (PharMingen) that recognizes both the 32 kDa pro-form and the 17 kDa active form of caspase-3 (Keane et al., 1997; Krajewska et al., 1997). Bound antibodies were visualized using a mouse anti-rabbit secondary antibody conjugated with horseradish peroxidase and detected by enhanced chemiluminescence according to the manufacturer's protocol (Amersham Phamacia Biotech, Piscataway, NJ, USA).
Quantitation of caspase-3 enzymatic activity
Sperm lysates were prepared as described above for immunoblot analysis. An aliquot of sperm lysate was diluted with a solution containing interleukin 1B converting enzyme (ICE) buffer and the fluorogenic substrate Ac-DEVD-afc (N-acetyl-aspartate-glutamate-valine-aspartate-AFC, 7-amino-4-trifluoromethyl coumarin) (Parvathenani et al., 1998). Fluorescent emission (excitation 400 nm and emission 505 nm) was measured after incubation for 45 min at 37°C. Blanks without sperm were evaluated to determine background fluorescence. Standards containing 0–500 pmol/l AFC were utilized to determine the amount of fluorochrome released. Fluorescence was measured at 
max = 505 nm using a SpectraMax Gemini XS (Molecular Devices, Sunnyvale, CA, USA). Caspase activity was expressed as pmol/min/mg protein. Apoptotic human neutrophils treated with 1 mmol/l cycloheximide were used as positive controls (Parvathenani et al., 1998). The level of sensitivity of this assay was determined to be 250 000 cells using standard curves of caspase activity in known numbers of neutrophils, Jurkat cells and HL-60 cells.
Annexin V binding assay
Translocation of PS to the outer leaflet of the plasma membrane was detected using the Annexin V Cy3.18 Apoptosis Detection Kit (Sigma) according to the manufacturer's instructions. At the onset of apoptosis, PS, normally found on the cytoplasmic face of the plasma membrane, translocates to the extracellular leaflet. Annexin V Cy3.18 (red fluorescence) binds to PS on the exterior surface of membranes of cells that are undergoing apoptosis. A second stain, 6-carboxyfluorescein diacetate (6-CFDA, green fluorescence), is used to assess viability and differentiate between apoptotic and necrotic cells. Using fluorescent microscopy, living cells stain only with 6-CFDA (live, annexin V-negative, green, normal cells). Necrotic cells stain only with annexin V Cy3.18 (red, dead cells). Early in apoptosis, cells stain with both annexin V Cy3.18 (red) and 6-CFDA (green), and are therefore green–red, live, annexin V-positive cells (Duru et al., 2000, 2001a,Duru et al., b).
Aliquots of sperm suspension were placed on poly-L-lysine-coated slides (Sigma) and incubated at room temperature before washing with binding buffer and incubation with the double label staining solution (annexin V Cy3.18 and 6-CFDA). Analysis of the samples was performed using epifluorescence microscopy as mentioned above. At least 100 sperm were assessed per slide over five random fields. The intra- and inter-observer variabilities were <6% for this technique (Duru et al., 2001a,b).
TUNEL assay
The DNA strand breaks were identified by TUNEL assay. An in-situ cell death detection kit, fluorescein (Boehringer Mannheim, Indianapolis, IN, USA) was used to detect DNA fragmented sperm with epifluorescent microscopy (see above) according to the manufacturer's recommended protocol. A positive control was prepared by incubating an aliquot of sperm with 1 mg/ml deoxyribonuclease I. For the negative control, no terminal transferase enzyme was added. At least 200 cells were analysed in 10 random fields for each slide. Each cell was classified as apoptotic (intense green nuclear fluorescence) or normal (no fluorescence). The intra- and inter-observer coefficients of variation were <7% for the technique (Duru et al., 2001a,b).
Experimental design
In these studies, we examined 25 patients and five donors. One ejaculate from each patient (n = 25 ejaculates) and ejaculates from the five donors (n = 9 ejaculates) were independently assayed. For each ejaculate, and after the separation of the individual semen fractions of high and low sperm motility, the following were assessed: motility parameters, morphology, TUNEL, annexin V binding and caspase activation.
Statistical analysis
The purified fractions with high and low sperm motility were compared with regard to all parameters by paired t-tests. Results of patients versus donors were compared by unpaired t-tests. Statistical relationships between parameters were assessed by Pearson's correlation coefficient in a correlation matrix. Results of dose- and time-dependent experiments were analysed using a mixed model analysis of variance. Overall, results were expressed as the mean ± SE. Statistical significance was set at P < 0.05.
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Table I
details morphology and motion parameters of the fractions with high and low sperm motility from patients and donors. Figure 1 shows immunostaining results with a monoclonal antibody to active, but not inactive, caspase-3 used to identify sperm with active caspase-3 in situ. Overall, no more than 5% of sperm showed positive immunostaining. The low motility fractions of patients and donors showed significantly higher levels of positively immunostained cells than the high motility fractions. Patients demonstrated higher levels of positively immunostained sperm than donors; however, the differences were not statistically significant. Figure 2A shows representative results of caspase-3 immunostaining in six sperm: one positive (see intense fluorescence in the midpiece) and five negative cells (one double-headed); Figure 2B depicts the same sperm as seen under bright field microscopy. In all positively identified cases, activated caspase-3 was localized exclusively to the sperm midpiece region.
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In a different approach, the presence of active caspase-3 was determined in sperm extracts using immunoblot analysis and a polyclonal antibody that recognizes both inactive and active caspase-3. Figure 3
shows results from representative patients and control donors. Inactive procaspase-3 was detected in all fractions of low sperm motility from patients and donors. Active caspase-3 was detected in a limited number of low motility Percoll fractions from patients and, in even fewer fractions, donors. Low levels of inactive procaspase-3 were detected with a larger variability in the high motility fractions from patients and donors; however, none of the high motility fractions exhibited active caspase-3.
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A third approach was used to determine the presence of active caspases in sperm extracts using the fluorogenic substrate DEVD-afc (Figure 4
). Since this substrate can be hydrolysed by several different caspases, including caspase-3, -6, -7 and -8, it provided a more generalized approach to include the possibility that caspases other than caspase-3 could be present in sperm, and allowed a quantifiable detection of caspase enzymatic activity. In the high motility sperm fractions, there was a significant difference, with a higher activity in patients than in donors. In the low motility fractions, there was also higher activity in patients than in donors, but the difference was not significant. Caspase enzymatic activity was much more readily detected in cycloheximide-treated human neutrophils, which served as positive controls.
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The caspase activity in the immunoblots and the enzymatic assays was not due to non-sperm cells because they were effectively removed before analysis by paramagnetic beds coated with CD45 antibodies. Furthermore, based on standard curves of caspase activity in known numbers of neutrophils, Jurkat cells and HL-60 cells, the levels of sensitivity for the enzymatic assay, which was more sensitive than immunoblot analyses, was ~250 000 cells, which could be readily detected by microscopy. Microscopic analysis did not identify cells other than sperm in any fractions analysed. The relatively low level of caspase activity in sperm is consistent with their small cytoplasmic volume and/or the likelihood that subpopulations of sperm do not exhibit elevated caspase activity.
To determine whether sperm exhibited caspase-dependent events that are well characterized in somatic cells, we evaluated annexin V binding as an early event in apoptosis and DNA fragmentation as a late apoptotic event. Annexin V binding results are shown in Figure 5. Patients had a significantly higher percentage of annexin V-positive, live sperm than donors in both the high and low motility fractions. In addition, the high motility fractions (in both patients and donors) had a significantly lower percentages of cells with PS on their outer membranes than the low motility fractions. Figure 6 shows representative patterns of sperm as observed with the annexin V binding assay: live cells without PS externalization (green, Figure 6A), dead cells (red, Figure 6B) and live cells with PS externalization (green-red, Figure 6C).
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TUNEL assay results are shown in Figure 7
. Patients and donor results were not significantly different with respect to DNA fragmentation. Both groups had significantly greater percentages of cells with DNA fragmentation (TUNEL-positive) in the low compared with the high motility fractions. Figure 8A shows a representative spermatozoon positive for TUNEL staining demonstrating DNA fragmentation and several negative sperm cells with no DNA fragmentation. Figure 8B depicts the same sperm as observed under phase contrast microscopy, and Figure 8C shows the same sperm using combined phase contrast and fluorescent light.
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In the high motility fractions, there was a significant negative correlation between motility and the percentage of annexin V-positive, live sperm (r = –0.59, P < 0.05) and caspase activity (r = –0.54, P < 0.05). Similar significant relationships were observed for the percentage of rapid sperm. A positive correlation (r = 0.5, P < 0.05) was detected between the annexin V binding and TUNEL results. In addition, there was significant positive correlation (r = 0.6, P < 0.05) between the TUNEL and caspase immunostaining results.
In the low motility fractions, there was a significant negative correlation between motility and the percentage of annexin V-positive, live sperm (r = –0.7, P < 0.05) and the percentage of sperm with DNA fragmentation (r = –0.65, P < 0.05). Similar significant relationships were observed for the percentage of rapid sperm. Significant positive correlations between caspase immunostaining and TUNEL results (r = 0.51, P < 0.05) and between caspase immunostaining and caspase enzymatic activity (r = 0.69, P < 0.05) were demonstrated. A significant positive correlation between annexin V binding (live cells with PS translocation) and TUNEL results (r = 0.6, P < 0.05) was also detected.
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Discussion |
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Caspase-dependent apoptosis is a well characterized and ubiquitous mechanism in eukaryotes for removing senescent, defective or unneeded somatic cells, but roles for caspases and apoptosis in ejaculated sperm are still in question. To address this issue, we analysed sperm fractions with high and low motility from healthy fertile donors and infertile patients for the presence of caspase activation as well as early and late caspase-dependent markers that have been extensively evaluated in somatic cells. To the best of our knowledge, this is the first unequivocal evidence for the presence of active caspases (including caspase-3) in ejaculated human sperm of both infertile patients and fertile donors. Here, we have presented evidence for the presence of active caspases by three different methods, including immunostaining for active caspase-3 in situ, immunoblotting with a caspase-3-specific antibody that identifies inactive as well as active caspase-3, and a more general in-vitro caspase enzymatic activity assay.
These studies also show that PS exposure on the outer membrane and DNA fragmentation, two of the well-characterized post-caspase events in somatic cells, are present in ejaculated sperm. Furthermore, the relative numbers of sperm that exhibited these markers were significantly negatively correlated with motility and numbers of rapid sperm in the low motility fractions. However, it is not clear whether the lower sperm motility and lower numbers of rapid sperm are the result or the cause of a high rate of apoptosis in these populations.
Activated caspase-3 was localized by immunostaining exclusively to the midpiece area of all positively stained sperm, suggesting that caspase-dependent apoptosis may be sequestered in a region where the mitochondria and remnants of the cytoplasmic droplet would be located in abnormal and/or immature sperm. This is consistent with the work of Blanco-Rodriguez and Martinez-Garcia who suggested that apoptosis is restricted to the cytoplasmic component of rat spermatids (Blanco-Rodriguez and Martinez-Garcia, 1999).
Several lines of evidence associated low sperm motility and poor morphology with the presence of high caspase, regardless of the method for caspase determination. Although no more than 5% of the sperm were positive for active caspase-3 in any given sample, the low motility fraction exhibited more active caspase-positive cells than the high motility fraction in both donors and patients. In addition, there were lower and more variable levels of procaspase-3 in high motility fractions of patients and donors, and a virtual absence of active caspase-3. Furthermore, higher levels of active caspase-3 (17 kDa) and the inactive pro-form (32 kDa) were present in the low motility compared with the high motility fractions from donors and patients.
Finally, using a quantitative caspase activity assay that identifies several caspases, including caspase-3, -6, -7 and -8, and ruling out caspase contributions from cells other than sperm, patients were shown to have higher caspase enzyme activity than donors in both the low and high motility fractions and donors had higher activity in the low than the high motility fraction. Some of this caspase activity is due to caspase-3, but other caspases may be detected by this method. This would be in agreement with Weil et al. who demonstrated that 8% of dead mouse sperm were stained by anti-caspase-3 antibodies using affinity-purified rabbit antibodies by an indirect immunofluorescence assay and postulated that mouse sperm have a death programme that may not depend on caspase-3 (Weil et al., 1998).
Regardless of the type of caspase isoform present in sperm, the levels of caspase activity found in (non-stimulated) ejaculated sperm were ~100-times lower than those found in cycloheximide-stimulated human blood neutrophils. Thus, it could be argued that all three assays suggest that caspase activation may be relatively uncommon in ejaculated sperm. On the other hand, not all sperm in the population are likely to exhibit active caspases and given that the cytoplasmic volume of sperm is small compared with neutrophils, it may be that in sperm caspase enzyme activity is more efficient, requiring low levels. In any event, the close correlation of high caspase and low motility suggests a possible association and that elevated caspase activity may be a marker of poor sperm quality.
We found that the degree of PS externalization and DNA fragmentation appeared to be relatively higher than the percentage of sperm that had positive immunostaining for active caspase-3. Importantly, there also appeared to be agreement between the percentage of sperm with cytoplasmic droplets and the percentage of caspase-3 immunostained cells. The lack of correlation between midpiece defects and caspase immunostaining may be due to the fact that the population of patients studied here did not suffer from severe teratozoospermia or oligoasthenoteratozoospermia. It will be interesting to examine infertile men presenting with severely amorphous sperm having significant midpiece defects and large cytoplasmic droplets. It is tempting to speculate that, in the sperm cells, caspase activation may have occurred earlier (at ejaculation or even during late spermiogenesis and/or epididymal storage or transit) and that it manifested as a high occurrence of apoptotic changes but low levels of active caspases as the mature sperm do not have efficient operative mechanisms for protein synthesis. In other words, there may be a temporal dissociation between caspase activation and the expression of cellular changes suggestive of apoptosis. Alternatively, triggering of PS externalization and DNA fragmentation could be due to activation of other caspases or cellular pathways.
While some apoptotic events are caspase-independent, caspase activation is believed to be a well defined point of no return for apoptosis progression in somatic cells, and a number of apoptotic events downstream of caspase activation have been characterized (Thornberry and Lazebnik, 1998). Therefore, we evaluated two well-known somatic cell caspase-dependent events, annexin V binding to PS on the outer leaflet of membrane, a relatively early apoptotic event, and DNA fragmentation, a relatively late apoptotic event. In both the donor and patient groups, annexin V binding and DNA fragmentation were greater in the low motility than in the high motility fractions. Annexin V binding, but not DNA fragmentation, was greater in patients than in donors for both fractions. Furthermore, positive correlations were observed with low motility fractions for caspase activation, annexin V binding and DNA fragmentation, suggesting a relationship between them.
However, as mentioned above, the frequency of annexin V binding and DNA fragmentation was higher than the percentage of cells that exhibited active caspase-3 by in-situ immunostaining. This suggests that if the two markers are caspase-dependent, caspase-3 may not be a primary determinant of either one of them and suggests the potential for other caspases to mediate these apoptotic events in ejaculated sperm. While the study showed that elevated caspase activity, PS exposure and DNA fragmentation occur in the same population of sperm, it did not clearly show that caspase activation causes PS exposure or DNA fragmentation. This also leaves open the possibility that sperm apoptosis may be caspase-independent, at least to some extent.
It is not clear if these apoptosis markers of sperm appeared before or after ejaculation in those sperm populations showing signs of immaturity or poor quality. It is tempting to speculate that they may appear after ejaculation because annexin V binding, an early apoptotic event, was more strongly correlated with low motility and low rapid sperm than DNA fragmentation, a late apoptotic event. Alternatively, these markers may indicate an abortive apoptosis mechanism that was interrupted at some stage of spermatogenesis causing the seminiferous tubule release of sperm with PS externalization and DNA fragmentation (Sakkas et al., 1999). It therefore remains unclear whether triggering of apoptosis occurs before or after the completion of spermatogenesis and whether defective or senescent sperm are typically eliminated by apoptosis before or after ejaculation.
It is known that DNA fragmentation, a consequence of apoptosis, occurs at the testicular level in spermatocytes and spermatids (Brinkworth et al., 1995; Callard et al., 1995; Lin et al., 1997; Sinha Hikim et al., 1998). On a theoretical basis, other apoptotic stimuli could trigger cell death following testicular sperm release, for example at the epididymal or other male ductal level; or apoptosis could be the result of post-ejaculatory events in the female tract or even upon in-vitro sperm incubation (Barroso et al., 2000). Apoptosis evolved, in part, to prevent the transmission of genetic defects to successive generations and this is as important in sperm, which generate new life, as it is to somatic cells, which maintain it. Therefore, it is highly likely that sperm may utilize similar somatic cell apoptosis mechanisms since apoptotic cell death is highly conserved and ubiquitous in eukaryotes.
In several studies, we and others have reported variable amounts of DNA fragmentation in the high and low motility fractions of ejaculates from infertile men and normozoospermic donors. In previous experiments, we found 1% DNA fragmentation in the purified fractions of highly motile sperm and 11% DNA fragmentation in the fractions of low motility using TUNEL (Barroso et al., 2000). In reasonable agreement with those results, Sun et al. used TUNEL and fluorescence-activated cell sorting to report that the percentage of sperm with DNA fragmentation was <4% after swim-up separation in the majority of samples of motile sperm from infertile men, but ranged from 5 to 40% in ~27% of the samples (Sun et al., 1997). Aitken et al. demonstrated that >30% of highly motile sperm (88% Percoll layer) from normozoospermic donors displayed DNA fragmentation using the comet assay (Aitken et al., 1998). Lopes et al. used TUNEL to show that the percentage of sperm with DNA fragmentation was <4% (ranging from 0 to 16%) in the majority of highly motile sperm after swim-up separation from infertile men (Lopes et al., 1998). Oosterhuis et al. concluded that 20% of sperm showed DNA fragmentation using TUNEL in ejaculated sperm of infertile men (Oosterhuis et al., 2000). Donnelly et al. showed that 40% of sperm from semen and 20% of sperm from the fractions with high sperm motility from infertile men had DNA fragmentation using TUNEL (Donnelly et al., 2000). The differences in the proportion of sperm with DNA fragmentation among all these studies may be due to the sperm samples analysed (donors versus different patient populations), the various sperm separation methods, and the different methods used to detect DNA fragmentation.
In our experiments, samples of live sperm from both high and low motility fractions from all ejaculates displayed some degree of staining with annexin V. PS, which is normally confined to the inner leaflet of the plasma membrane, may act as a membrane `flag' on apoptotic cells (Fadok et al., 1992). Redistribution of PS possibly affects membrane stability and charge, results in membrane asymmetry, disruption of channels, and receptor activities. Since it is known that membrane phospholipid changes occur during capacitation and the onset of the acrosome reaction, more studies are needed to definitely establish whether annexin V binding results represent true apoptotic changes in sperm.
In general, all apoptosis markers (active caspase, annexin V binding and DNA fragmentation) were higher in infertile patients than in donors and greater in the low versus the high motility fractions. However, statistical significance was often marginal in the patient population, which exhibited greater variability as indicated by larger standard errors when compared with donors. Thus, it is possible that subsets of patients exhibit apoptosis markers that are diagnostic for infertility, but these are `diluted out' by other patients who are infertile for reasons unrelated to apoptosis or are mistakenly diagnosed. Further investigation and patient classification may identify relationships between apoptosis and infertility that define a type of reproductive failure that could be modulated therapeutically.
In conclusion, sperm fractions with higher frequencies of caspase activation as recorded by immunostaining were correlated with a higher degree of DNA fragmentation in infertile patients and in fractions with low motility, possibly indicating caspase-dependent apoptosis. The exclusive immunostaining detection of active caspase-3 to the sperm midpiece region suggests that a caspase-dependent apoptotic mechanism may originate in the cytoplasmic droplet or within mitochondria. However, it cannot be ruled out that caspase-independent mechanisms may be operative in human sperm or that sperm do not utilize the same mechanisms for cell death as somatic cells. The possibility also exists that some of the phenomena observed (such as externalization of PS) may represent functional changes that sperm undergo both under in-vivo and in-vitro conditions. However, we suggest that small numbers of sperm, particularly immature and/or abnormal cells found in the low motility fractions, may have an operative caspase-dependent programmed cell death in their final stages of development. Roles for caspase-dependent and -independent apoptosis mechanisms in post-ejaculated human sperm clearly require further analysis. Understanding of these mechanisms may provide the means to manipulate apoptosis for therapeutic gain.
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Acknowledgements |
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We are thankful to Dr P.Kolm, Bio-statistician, Portsmouth Naval Hospital, Portsmouth, Virginia, for his assistance in data analysis. This study was supported by the Commonwealth of Virginia Health Research Board and by the Jeffress Memorial Trust.
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4 To whom correspondence should be addressed at The Jones Institute for Reproductive Medicine, 601 Colley Avenue, Norfolk, VA 23507, USA. E-mail: oehninsc{at}evms.edu
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References |
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Submitted on November 23, 2001; resubmitted on April 4, 2002; accepted on July 19, 2002.
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