Investigation on the Origin of Sperm DNA Fragmentation: Role of Apoptosis, Immaturity and Oxidative Stress
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Sperm DNA fragmentation (sDF) represents a threat to male fertility, human reproduction and the health of the offspring. The causes of sDF are still unclear, even if apoptosis, oxidative assault and defects in chromatin maturation are hypothesized. Using multicolor flow cytometry and sperm sorting, we challenged the three hypothesized mechanisms by simultaneously evaluating sDF and signs of oxidative damage (8-hydroxy, 2′-deoxyguanosine (8-OHdG) and malondialdehyde (MDA)), apoptosis (caspase activity and cleaved poly(ADP-ribose) polymerase (cPARP)) and sperm immaturity (creatine phosphokinase (CK) and excess of residual histones). Active caspases and c-PARP were concomitant with sDF in a high percentage of spermatozoa (82.6% ± 9.1% and 53.5% ± 16.4%, respectively). Excess of residual histones was significantly higher in DNA-fragmented sperm versus sperm without DNA fragmentation (74.8% ± 17.5% and 37.3% ± 16.6%, respectively, p < 0.005), and largely concomitant with active caspases. Conversely, oxidative damage was scarcely concomitant with sDF in the total sperm population, at variance with live sperm, where 8-OHdG and MDA were clearly associated to sDF. In addition, most live cells with active caspase also showed 8-OHdG, suggesting activation of apoptotic pathways in oxidative-injured live cells. This is the first investigation on the origin of sDF directly evaluating the simultaneous presence of the signs of the hypothesized mechanisms with DNA breaks at the single cell level. The results indicate that the main pathway leading to sperm DNA breaks is a process of apoptosis, likely triggered by an impairment of chromatin maturation in the testis and by oxidative stress during the transit in the male genital tract. These findings are highly relevant for clinical studies on the effects of drugs on sDF and oxidative stress in infertile men and for the development of new therapeutic strategies.
In the last two decades we have been aware that in human ejaculates there can be high percentages of sperm with DNA fragmentation, representing a threat for male fertility, human reproduction and the health of the offspring. In addition, in the era of assisted reproduction techniques (ARTs) which bypass many, if not all, natural barriers to fecundation, the risk that sperm with unresolved DNA damage can fertilize an oocyte (1) appears increased, thus raising further concerns on the presence of DNA breaks in the sperm genome. The first reports on sperm DNA fragmentation (sDF) date back to the late 1980s (2) and early 1990s (3) and, since then, the biological and clinical meanings of this type of sperm damage have been investigated extensively and several techniques to reveal it have been developed (4). However, the causes and the sites of origin of sDF have not been completely clarified and, still, we are dealing with hypotheses and theories. Clearly, the knowledge of the mechanisms responsible for this type of sperm damage is pivotal for the development of effective treatments to prevent the onset of sDF in infertile men.
Besides well-known external inducers of sDF, including chemotherapy (5), environmental toxicants (6) and the presence of leukocytes in semen (7), three main mechanisms have been proposed to explain the genesis of sDF. According to one of these proposed mechanisms, the DNA nicks occurring to promote the remodeling of sperm chromatin are not completely repaired due to an impairment of the sperm maturation process (8,9). sDF also could reflect a DNA cleavage produced by a process of apoptosis first triggered and later interrupted in the testis (that is, abortive apoptosis) (10) or occurring after spermiation (11,12). Finally, sperm DNA breaks could be provoked by attacks of free radicals, including reactive oxygen species (ROS) (13), acting both in testis and in posttesticular sites (14,15). It is anticipated that these proposed mechanisms are not alternative, but can concur in generating the sperm DNA damage. Indeed, besides the occurrence of persistent DNA nicks, an impairment in chromatin maturation could produce poorly compacted nuclei, which are more vulnerable to oxidative assault (13). Similarly, ROS could break the DNA backbone directly, but also act as triggers of apoptotic pathways ending in caspases and apoptotic nucleases activation, as happens in somatic cells (16). The above hypotheses are supported by indirect studies showing that infertile/subfertile subjects who are known to have increased levels of sDF show higher degrees of cell immaturity (17) or of apoptotic features (18, 19, 20) or of oxidative stress (21,22) in their ejaculates and by correlative studies reporting associations between the levels of sDF and signs of impaired chromatin maturation, (23,24) or apoptotic traits (25,26) or evidence of oxidative stress (27,28). However, these approaches detect the markers of the hypothesized mechanisms in different semen aliquots from those used to reveal sDF and the occurrence of statistical correlations does not necessarily imply a cause-effect relationship. Only in a few studies (24,28) has a direct approach been used to study the mechanisms responsible for sDF by double-staining sperm for sDF and chromatin immaturity markers. In such studies (24,28), the latter are strictly linked to sperm DNA breaks as observed in a small number of cells by microscopy. Sakkas et al. (29) employed flow cytometry to address the relation between sDF and apoptotic proteins, failing, however, in revealing a link between sperm DNA breaks and apoptosis, possibly because of the inclusion in the fluorescence analyses of semen apoptotic bodies, scarcely TUNEL-labeled (30) but highly expressing apoptotic features (31).
In the present study, we challenged the hypothesized mechanisms and their relative contribution to the genesis of sDF by directly investigating the presence of signs of apoptosis, chromatin immaturity and oxidative stress in a large number of sperm with and without DNA fragmentation. To this aim, we used multicolor flow cytometry to simultaneously detect sDF and signs of oxidative damage (8-hydroxy, 2′-deoxyguanosine [8-OHdG] and malondialdehyde [MDA]), of apoptosis (active caspases and cleaved poly[ADP-ribose] polymerase [cPARP]) and of sperm immaturity (creatine phosphokinase [CK]) (32,33). The association between sDF and chromatin immaturity also was investigated by aniline blue (AB) staining in fractions of sperm with and without sDF, separated by fluorescence-activated cell sorting. To our knowledge, this is the first study on the origin of sDF where the concomitance of this sperm damage with the possible causes responsible for it was investigated by a high throughput strategy.
Materials and Methods
Human tubal fluid (HTF) medium was purchased from Celbio (Milan, Italy). Bovine serum albumin (BSA) was purchased from ICN Biomedicals (Irvine, California, USA). The primary antibodies used in the study were: monoclonal mouse antibodies anti 8-OHdG, 15A3 (Santa Cruz Biotechnology, Santa Cruz, CA, USA) and anti MDA, clone 1F83 (JaICA, Haruoka, Japan); CKB antibody (N-term), purified rabbit polyclonal antibody (Abgent, San Diego CA, USA). Antibodies for secondary detection were: the goat anti-rabbit IgG (H + L chain specific), fluorescein (FITC) conjugate (Southern Biotech, Birmingham, AL, USA); the goat anti-mouse IgG-FITC and the sheep anti-mouse IgG (whole molecule) F(ab’)2 fragment-R-phycoerythrin (R-PE) (Sigma-Aldrich, St. Louis, Missouri, USA). Mouse IgG2a isotype control antibody was purchased from Exbio (Praha, Czech Republic). Anti-PARP CSSA FITC, apoptosis detection kit and Vybrant FAM Caspase-3 and −7 Assay Kit were purchased from Life Technologies (Paisley, UK [Thermo Fisher Scientific Inc., Waltham, MA, USA]).
The study has been approved by the Local Ethical Committee of the Azienda Ospedaliera e Universitaria (AOUC) Careggi, and informed written consensus has been obtained from the recruited patients.
Semen Sample Collection
Semen samples were consecutively collected according to WHO criteria (34) from men undergoing routine semen analysis as part of testing of couples with fertility problems in the Andrology Laboratory of the University of Florence. Subjects undergoing drug therapies were excluded from the study as well as semen samples where leukocytes exceeded 1 million/mL. Occurrence of leukocytes was assessed by counting all round cells in the Neubauer chamber and then distinguishing leukocytes from germ cells after differential quick staining of the sample. For the experiments of the study, semen samples were processed individually and different semen samples were used for each marker. Overall, we recruited 92 subjects (average age: 34.8 ± 7.8 years) showing the following average semen parameters values: normal morphology, 6.2 ± 4.8%; total motility, 60.5 ± 20.9%; progressive motility, 56.7 ± 14.5%; concentration, 70.6 ± 41.8 millions/mL; total number/ejaculate, 288.1 ± 137.5 millions. After liquefaction (30 min following collection according to ), semen samples were washed twice with HTF medium and, treated by dithiothreitol (DTT, 2 mmol/L, 45 min at room temperature) (35), washed again twice with the same medium and, unless otherwise indicated, fixed by 500 µL of 4% paraformaldehyde in phosphate buffered saline (PBS), pH 7.4, for 30 min at room temperature.
For experiments of treatment of sperm with H2O2, semen samples were washed twice and split into two aliquots that were incubated in the medium containing or not 5 mmol/L H2O2 for 2 h at 37°C (36). After washing, samples were then treated by DTT and fixed as described above.
For experiments in live sperm, after washing with HTF medium, fresh semen samples were incubated for 1 h at room temperature, in the dark, in 500 µL of PBS with Live/Dead Fixable Far Red Dead Cell Stain Kit (L10120, diluted 1:10 000; Life Technologies [Thermo Fisher Scientific]), and treated with DTT and fixed as described above. L10120 is able to bind dead cells and the labeling is stably kept in the cells after sample fixation and permeabilization (35,37,38).
For labeling sDF, samples were processed by terminal deoxynucleotidyl transferase (TdT)-mediated fluorescein-dUTP nick end labeling (TUNEL) as described elsewhere (39). Briefly, fixed sperm were washed by PBS/BSA 1% twice and permeabilized with 0.1% Triton X-100 in 100 µL of 0.1% sodium citrate for 4 min in ice. After washing two times, the labeling reaction was performed by incubating sperm in 50 µL of labeling solution (supplied with the In Situ Cell Death Detection Kit, fluorescein, Roche Molecular Biochemicals, Milan, Italy) containing the TdT enzyme, for 1 h at 37°C in the dark. Finally, samples were washed twice, resuspended in PBS, stained with PI (propidium iodide, 0.75 µg/mL) and incubated in the dark for 15 min at room temperature. For each test sample, a negative control was also prepared by omitting TdT.
Positive controls for TUNEL were prepared: (i) by incubating sperm with 2 mmol/L H2O2 in HTF medium (3 h, 37°C) before processing samples for TUNEL; (ii) by treating sperm with DNAse I (Pharmacia Biotech Italia, Milan, Italy), 2 IU for 20 min at 37°C, before the labeling reaction.
In preliminary experiments, we assessed whether the pretreatment of samples with DTT increased the sensitivity of TUNEL. In agreement with a previous study (35), we found that DTT treatment increased the percentage of sDF from 31.8 ± 15.8% to 37.9 ± 19.0% (p < 0.01, n = 10).
Simultaneous Detection of Oxidative Signs (8-OHdG and MDA) and sDF
Fixed sperm (20 × 106) were washed (twice with 1% normal goat serum [NGS]-PBS) and split into two aliquots. For 8-OHdG detection, the two aliquots were incubated in 100 µL of 0.1% sodium citrate/0.1% Triton X-100 containing 2 µg/mL anti-8-OHdG antibody 15A3 (test sample) or 2 µg/mL mouse IgG2a (isotype control) for 1 h at 37°C (40). For MDA detection, the aliquots were first permeabilized in 100 µL of 0.1% sodium citrate/0.2% Triton X-100 (30 min at room temperature) and then incubated in 100 µL of 1% NGS-PBS containing 2 µg/mL of the antibody against MDA (test sample) or 2 µg/mL mouse IgG2a (isotype control) for 1 h at 37°C (41). After incubating with the antibodies and washing twice with 1% NGS-PBS, sperm were incubated in the dark (1 h at room temperature) with FITC-conjugated goat anti-mouse IgG (dilution 1:100 in 100 µL 1% NGS-PBS). Then, both the test sample and the isotype control were split again into two aliquots and each aliquot was processed for TUNEL assay as described above. However, to avoid the overlapping between fluorescence signals emission, we used TMR (tetramethylrhodamine)-conjugated dUTPs (39) supplied with the In Situ Cell Death Detection Kit, TMR (Roche Molecular Biochemicals). For similar reasons, nuclear staining was performed by DAPI (1 µg/mL for 15 min in the dark at room temperature) instead of PI. DAPI staining is able to discriminate between M540 bodies and sperm and, within the latter, between brighter and dimmer populations (see Results section) similarly to PI staining. Indeed, in preliminary experiments (n = 5 semen samples) we found that the percentages of sperm within the flame-shaped region in forward scatter/side scatter (FSC/SSC) dot plots (see below) and of the two populations versus total sperm do not change by staining with DAPI and PI (data not shown).
In some experiments (n = 3), we assessed the reproducibility of the above procedures by processing semen samples in duplicate and comparing the two measures. In the case of 8-OHdG/TUNEL, we found an average coefficient of variation (CV) of 4.9% ± 5.0% for TUNEL and of 2.1% ± 1.0% for 8-OHdG. In the case of MDA/TUNEL, the average CV of the measures was 7.9% ± 3.9% for TUNEL and 19.2% ± 15.1% for MDA.
We also verified whether the double-labeling procedure affected the measures of each parameter as if separately determined. Regarding TUNEL/MDA detection, the average CVs of the measures obtained by double and single labeling were 12.1% ± 1.0%, (n = 3) for TUNEL and 20.0% ± 5.5%, (n = 3) for MDA. Similarly, the measures of sDF as assessed by double (TUNEL/8-OHdG) and single labeling were fairly consistent (average CVs = 7.9% ± 1.5% and 4.9% ± 5.4%, respectively, for TUNEL and 8-OHdG, n = 3).
Simultaneous Detection of CK and sDF
For CK detection (33), fixed sperm (20 × 106) were washed (twice with 1% NGS-PBS) and permeabilized in 100 µL of 0.1% sodium citrate/0.1% Triton X-100 for 4 min in ice. After washing with 1% NGS/PBS, sperm were blocked for 20 min in 5% NGS in PBS and washed again. Then the samples were split into two aliquots and incubated in 100 µL of 1% NGS-PBS containing 5 µg/mL of the antibody against CK (test sample) or 5 µg/mL of anti-rabbit serum (negative control) for 1 h at room temperature. After washing twice with 1% NGS-PBS, sperm were incubated in the dark (1 h at room temperature) with the FITC-conjugated goat anti-rabbit IgG (1:100 in 100 µL of 1% NGS/PBS). Finally, samples were washed twice and both the sample test and the negative control were further split into two aliquots and each aliquot was processed for TUNEL assay as described above, using TMR-dUTPs for the labeling of DNA breaks and DAPI for nuclear staining.
In some experiments (n = 3), we assessed the reproducibility of the above procedure by processing semen samples in duplicate and comparing the two measures. We found an average CV of 12.4% ± 5.1% for TUNEL and of 14.9% ± 19.7% for CK expression.
We also verified whether the double-labeling procedure affected the measures of each parameter as if separately determined. The average CVs of the measures obtained by double TUNEL/CK labeling and single labeling were 14.9% ± 6.8% (n = 3) for TUNEL and 7.1% ± 1.6% (n = 3) for CK.
Simultaneous Detection of cPARP and sDF
For detection of cPARP (12), we used the FITC-conjugated anti-PARP cleavage site-specific antibody (CSSA) kit, following the instructions of the manufacturer, with slight modifications. Briefly, washed sperm samples (15 × 106) were fixed in 1 mL IC Fix buffer for 20 min at 4°C and, after washing twice and resuspension in PBS/BSA1%, split into two aliquots for negative control and test sample. In the latter, 10 µL of the antibody anti-cPARP were added and then the aliquots were incubated for 30 min at room temperature. After washing, the sample test and the negative control were split into two aliquots and each aliquot was processed for TUNEL assay as described above using TMR-dUTPs for the labeling of DNA breaks and DAPI for nuclear staining.
In some experiments (n = 3), we assessed the reproducibility of the above procedure by processing semen samples in duplicate and comparing the two measures. We found that the average CV of the measures was 12.1% ± 3.9% for TUNEL and 15.5% ± 10.2% for cPARP.
We also verified whether the double labeling procedure affected the measures of each parameter as if separately determined. The average CVs of the measures obtained by double TUNEL/PARP labeling and single labeling were 3.5% ± 2.9% (n = 3) for TUNEL and 6.3% ± 3.3% (n = 3) for PARP.
Simultaneous Detection of Active Caspases and sDF
Caspases were evaluated by Vybrant FAM Caspase-3 and −7 Assay Kit which detects active caspases 3 and 7 by using the FAM-DEVD-FMK reagent (FLICA), a fluorescent-labeled inhibitor of such enzymes that covalently binds the enzymatic reactive center of the activated caspases (31). Briefly, sperm samples (15 × 106) were resuspended in HTF medium and split into two aliquots for the negative control and the test sample. In the latter, 10 µL of 30× FLICA working solution was added, then aliquots were incubated for 1 h at 37°C in the dark. After two washes with wash buffer 1× (included in the kit mentioned above), samples were fixed by adding 40 µL of 10% formaldehyde solution (included in the kit mentioned above) for 10 min at room temperature. Then, sperm samples were washed again twice and both negative control and test sample were split again into two aliquots that were processed for TUNEL assay as described above, using TMR-dUTPs for the labeling reaction and DAPI for nuclear staining.
In selected experiments we investigated the concomitance of 8-OHdG and active caspases in live sperm. To this end, after labeling with L10120, sperm samples were processed for detection of caspase activity by the above procedure. Then, both the negative control and test sample were split into two aliquots that, in turn, were labeled for 8-OHdG detection as described above, but using an antibody conjugated with R-PE for secondary detection to avoid fluorescence of 8-OHdG that overlapped that of FLICA.
In some experiments (n = 3), we assessed the reproducibility of the above procedure by processing semen samples in duplicate and comparing the two measures. We found that the average CVs of the measures were 4.5% ± 5.5% for TUNEL and 4.7% ± 3.8% for activated caspases.
We also verified whether the double-labeling procedure affected the measures of each parameter as though separately determined. The average CVs of the measures obtained by double TUNEL/FLICA labeling and single labeling were 20.2% ± 2.8% (n = 3) for TUNEL and 26.3% ± 1.7%, (n = 3) for FLICA.
Fluorescence-Activated Cell Sorting
Fluorescence-activated cell sorting was used to separate TUNEL-positive from TUNEL-negative sperm. Samples containing double-labeled PI/TUNEL sperm were adjusted at concentration of 107cells/mL, filtered by 50-µm Syringe Filcons and immediately sorted by the BD FACSAria II cell-sorting system equipped with a FACSSort fluid sorting module. For sperm sorting, we used the followings settings: laser power, 13 mW; nozzle, 70 µm; sort setup, low; sheath pressure, 34.50 psi; frequency, 60.0 kHz; flow rate, 1–3 µL/min (maximum of 7,000 events/second); and precision, 0160. After gating to exclude large cells, debris and other semen interference (39), as described above, we used the negative control (omitted TdT) as reference for drawing two regions around the TUNEL negative and positive sperm. After sorting, we acquired the sorted fractions again by flow cytometry to check the purity of sperm with and without DNA fragmentation.
Staining with Aniline Blue (AB)
To evaluate sperm chromatin immaturity, we performed AB labeling (24,42). Briefly, after sperm sorting as described above, 100,000 sperm were smeared on slide, air dried and then stained with 5% aqueous AB (Sigma-Aldrich) mixed with 4% acetic acid (pH 3.5) for 5 min (42) at room temperature. Two hundred sperm were analyzed on each slide under a light microscope (Leica DM LS, Leica, Wetzlar, Germany). Sperm showing dark-blue staining were considered as AB positive, whereas those stained only weakly or not at all were considered as AB negative (43). After determining the percentage of fragmented sperm in presorted samples, the overall percentage of AB staining in such samples was calculated by (% AB staining in fragmented sperm × % fragmented sperm)/100 + (% AB staining in non-fragmented sperm × % nonfragmented sperm)/100.
Sperm labeled for sDF and for 8-OHdG, MDA, CK, active caspases or cPARP as described above, were spread onto slides and mounted with fluoromount aqueous mounting medium (Sigma Aldrich). Green and red fluorescence was examined using a fluorescence microscope (Axiolab A1 FL, Carl Zeiss, Milan, Italy), equipped with Filter set 15 and 44 by an oil immersion 100× magnification objective.
All variables were checked for normal distribution by the Kolgomov-Smirnov one-sample test. Since all variables resulted as normally distributed, results are expressed as mean ± SD and analysis of variance and the Student t test (paired and unpaired data) were used to assess statistically significant differences between the compared groups. A p value of 0.05 was considered as statistically significant. Coefficients of variation were calculated by (SD/mean) × 100. All statistical analyses were carried out using MicroCal Origin software, 6.1 version (MicroCal Software Inc., Northampton, MA, USA) except for evaluation of normal distribution, which was obtained by using SPSS software, 20 version, for Windows (SPSS [IBM, Armonk, NY, USA]).
All supplementary materials are available online at https://doi.org/www.molmed.org.
Multicolor Flow Cytometry
To study the simultaneous occurrence of apoptosis, immaturity and oxidative damage in sperm with sDF, we used flow cytometry to evaluate specific markers of each of the three processes in sperm samples stained by TUNEL and DAPI. Supplementary Figure 1 reports a flow chart depicting the experimental sets of the study. The staining with DAPI (Figure 1B) is necessary to define the sperm population (44) within the FSC/SSC region (Figure 1A), containing both sperm and semen apoptotic bodies lacking chromatin (31). Nuclear staining also unveils the occurrence of two sperm populations, named brighter and dimmer populations, based on a different intensity of such staining (44) (see Figure 1B). Brighter population represents the majority of semen sperm showing a normal DNA content (42), albeit fragmented in a variable percentage (44), whereas dimmer population represents roughly 15% of sperm and is totally composed by DNA-fragmented (44) and dead (45) cells, including those with large loss of chromatin material (41). Hence, data presented here, albeit referred to the total sperm population (region R1 in Figure 1B) unless otherwise indicated, also were determined by excluding dimmer sperm (that is, in the only brighter population, region R2 in Figure 1B).
Association of sDF with Sperm Immaturity
Association of sDF with Apoptosis: Caspase Activation and cPARP
Association of sDF with Oxidative Damage: 8-OHdG and MDA
The association between DNA breakage and oxidative insult was studied further by detecting another marker, the product of lipid peroxidation MDA, by using an immunofluorescent technique. The simultaneous staining for TUNEL and MDA showed, on average, a percentage of 17.7 ± 7.9 sperm (n = 11) with MDA and depicted a pattern of association of the two parameters similar to that with 8OHdG (compare Figure 6A and 6C). Indeed, the concomitance between expression of MDA and sDF was found in a small percentage of sperm, with most damaged sperm exhibiting sDF and MDA alternatively (Figure 6C and Supplementary Table 1). The incidence of sperm with MDA among those fragmented and non-fragmented was not statistically different (Figure 6D, left panel). However, as for 8OHdG (Figure 6B, right panel) the link between the membrane oxidative damage and DNA breaks became evident when we considered only the brighter sperm (Figure 6D, right panel). MDA was mostly localized in the midpiece of sperm (Figure 6E, left panel) (52).
Association of sDF with 8-OHdG, MDA and Active Caspases in Live Sperm
The above results suggest that oxidative and apoptotic signs can be concomitant in live sperm. To investigate this possibility, we simultaneously evaluated 8-OHdG and caspase activity in live sperm in three semen samples. As shown in Figure 7D, whereas most oxidized sperm did not show the apoptotic enzymes, most caspase-positive sperm also exhibited the 8-OHdG.
Understanding the causes of sDF is a goal sought over the last two decades, given the crucial importance of DNA integrity of spermatozoa for human reproduction and the health of the offspring. Using mostly multicolor flow cytometry analyses, we obtained postejaculation snapshots of how apoptosis-oxidation-immaturity (the three hypothesized mechanisms generating sDF) are distributed in total and live sperm with or without sDF, from which we can infer the pathways leading to such damage. Based on our results, the following conclusions can be drawn: (i) apoptosis is the main pathway leading to DNA breakage in sperm; (ii) chromatin immaturity induces sDF through activation of an apoptotic pathway; and (iii) oxidative attack appears to act after spermiation, occurring mostly in live sperm.
The finding that nearly all sperm with sDF also show the activity of the effector caspases 3 and 7 strongly indicates a key role of apoptosis in generating the bulk of sperm DNA breaks, consistent with the reported correlations between the levels of DNA breakage and of apoptotic traits in sperm (25,26). In addition, the causative relation between caspase activation and sDF also is suggested by the presence of a certain percentage of cells with the apoptotic enzymes but without DNA breaks, consistent with the idea that the activation of caspases precedes the beginning of sDF (46). The conclusion that apoptosis is the main mechanism responsible for sDF in the ejaculate was reinforced by the results obtained with cPARP, a different apoptotic marker (47,53), distributed among fragmented and nonfragmented sperm similarly to caspases.
The trigger of the apoptotic pathways can occur both in germ cells during spermatogenesis or after spermiation or both. Indeed, caspase activation and PARP cleavage have been observed both in the testis and in ejaculated sperm in response to apoptotic stimuli in vitro (12,50). The high percentage of sperm showing the simultaneous presence of sDF and both AB staining and active caspases or cPARP (present study), indicates that a large fraction of sperm concomitantly shows apoptotic traits and incomplete protamination, suggesting that apoptosis is triggered mostly in the testis. This conclusion is further supported by the presence of apoptotic bodies of testicular origin in semen of subfertile patients (20,31). However, the finding that the amount of sDF is greater in ejaculated sperm than in sperm extracted from the testis (14,15) indicates that apoptotic stimuli also can occur during transit in the male genital tracts. The fact that a fraction of live sperm also shows DNA fragmentation associated with active caspases is another indication that the onset of apoptosis may occur following release from the testis (see below for further discussion on this point).
Oxidative attack is the third main mechanism hypothesized to induce sDF. We found that the occurrence of 8-OHdG and MDA, hallmarks of oxidative stress, scarcely mirrored the presence of sDF. A statistically significant association between oxidation and breakage in DNA was found only in brighter sperm, that is, excluding dead and DNA-fragmented dimmer sperm (44,45), suggesting that the role of oxidation in generating sDF might be masked by the high amounts of DNA-fragmented, dead sperm present in the ejaculates of subfertile men (35). This prompted us to investigate the mechanisms of sDF in only the viable sperm fraction of the ejaculate, where DNA damage is still ongoing. In live sperm, the frequency of the copresence of 8-OHdG and MDA with TUNEL positivity was much greater, highlighting a causative role of oxidation. The finding that caspase activation is present in fragmented live sperm with a similar frequency of 8-OHdG and MDA, and that most apoptotic sperm also show the oxidized base, supports the idea that ROS could produce DNA breaks through apoptotic routes (Figure 8) rather than by directly breaking the phosphodiester backbone. This is supported also by the large presence of cells with 8-OHdG but without DNA breaks and is consistent with studies reporting that ROS induce apoptotic pathways in mammalian spermatozoa during in vitro incubation (50,62, 63, 64).
Overall, our results depict a scenario (Figure 8) where apoptosis, either if primed in testis or after spermiation, is the main causative mechanism of sDF. In the testis, the impairment of chromatin maturation appears to be a relevant apoptotic stimulus, leading to sDF and/or cell death, consistent with the idea that apoptotic germ cells may escape phagocytosis by Sertoli cells, in postmeiotic phases (65). Oxidative stress does not seem to play a relevant role in producing sDF in testis, as also suggested by the scarce occurrence of 8-OHdG in testicular tissue (66), possibly due to the presence of efficient DNA repair systems (67). After spermiation, during the transit in the male genital tracts, oxygen species become relevant in triggering apoptosis in live cells (Figure 8), but seem ineffective toward dead/dying cells primed to apoptosis in testis (52) explaining why, in the bulk of ejaculated sperm where most DNA-fragmented cells are dead (35), the oxidative signs are scarcely associated to sDF.
Recently, it has been proposed that TUNEL is able to detect only sDF generated in peri/post mortem sperm, at the end of processes of destruction and after the induction of a self-perpetuating ROS production and activation of caspases by oxidative stress (7,13,36,68,69). According to this hypothesis, sperm DNA is damaged only by ROS which generate oxidized adducts to DNA, including 8-HdG. However, if DNA-fragmented sperm were the final step of oxidative-injured cells, a large concomitance between 8-OHdG and MDA with DNA breaks should be observed, contrary to what this study found. In addition, we also found DNA breakage in a consistent percentage of live sperm, further questioning the interpretation of TUNEL positivity as a facet of moribund/dead sperm (13,36).
To the best of our knowledge, this is the first study investigating all the main mechanisms hypothesized for the origin of sDF by directly verifying the signs of such mechanisms in DNA-fragmented sperm. One limitation of the study is that the conclusions are mainly drawn on descriptive data, contrary to in vitro studies more suitable to trace cause-effect relationships between apoptotic/oxidative insults and sDF. However, it must be noted that in vitro-prepared sperm are scarcely representative of in vivo conditions and, as a result, are more vulnerable to oxidative assault because they are deprived of the antioxidant defenses of seminal plasma (70). In addition, in vitro investigations cannot address the involvement of mechanisms thought to be triggered in the testis, such as apoptosis and sperm chromatin maturation defects, because these mechanisms are acting in cells maturing in the testis and during epididymal transit. Finally, it has to be considered that during in vitro incubation, sperm chromatin might undergo changes in accessibility of the tools to reveal sDF due to the removal of zinc from chromatin (71,72). Such events could affect chromatin stainability both in ex vivo study (like ours) and (with a greater extent) during in vitro incubations.
In conclusion, our results indicate that the main pathway leading to sperm DNA breaks is a process of apoptosis triggered by testicular conditions and by oxidative stress during the transit in the male genital tract. The clarification of the mechanisms leading to DNA breaks may help to better focus studies aimed at evaluating the effect of drugs for male infertility (for instance, the effect of antioxidants should be evaluated in live sperm) and open new therapeutic perspectives for the treatment of the infertile men.
The authors declare they have no competing interests as defined by Molecular Medicine, or other interests that might be perceived to influence the results and discussion reported in this paper.
We are grateful to D Manganaro (Becton Dickinson, Milan, Italy) for precious technical assistance in the experiments of sorting spermatozoa. We also thank E Filimberti, S Degl’Innocenti and MG Fino (Azienda Ospedaliera-Universitaria Careggi), for evaluation of semen parameters. This study was supported by Regione Toscana (grant to G Forti), Ministry of Education and Scientific Research (PRIN 2009 project to E Baldi and FIRB project to S Marchiani).
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