Molecular Human Reproduction, Vol. 8, No. 3, 213-220,
March 2002
© 2002 European Society of Human Reproduction and Embryology
Testis and spermatogenesis |
Fas expression correlates with human germ cell degeneration in meiotic and post-meiotic arrest of spermatogenesis
1 Department of Internal Medicine, Andrology Unit, 2 Statistics and 3 Department of Experimental Medicine, Endocrinology Unit, University of L'Aquila, Italy
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Abstract |
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Degeneration of human male germ cells was analysed by means of light (LM) and transmission electron (TEM) microscopy. The frequency of degenerating cells was correlated with that of Fas-expressing germ cells in human testes with normal spermatogenesis (n = 10), complete early maturation arrest (EMA) (n = 10) or incomplete late maturation arrest (LMA; n = 10) of spermatogenesis. LM analysis of testis sections with normal spermatogenesis indicated that degenerating germ cells were localized in the adluminal compartment of the seminiferous epithelium. TEM showed that apoptotic cells were mostly primary spermatocytes and, to a lesser extent, round or early elongating spermatids. Apoptotic germ cells appeared to be eliminated either in the seminiferous lumen or by Sertoli cell phagocytosis. An increased number of degenerating cells was observed in testes with LMA as compared with normal testes and testes with EMA of spermatogenesis (P < 0.001, Wilcoxon's rank sum test). Comparison of these results with those obtained from immunohistochemistry experiments demonstrated a tight correlation between the number of apoptotic cells and the number of Fas-expressing germ cells (P = 0.001, Spearman's rank = 0.69). These findings suggest that altered meiotic and post-meiotic germ cell maturation might be associated with an up-regulation of Fas gene expression capable of triggering apoptotic elimination of defective germ cells.
apoptosis/Fas/immunohistochemistry/spermatogenesis/ultrastructure
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Introduction |
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Spermatogenesis is a dynamic process in which a cohort of undifferentiated diploid cells (spermatogonial stem cells) proceed throughout a sequence of mitotic and meiotic cell divisions and differentiation to generate spermatozoa (De Martino et al., 1989
). Although this process produces a remarkable number of sperm—200x106 per day in humans—up to 75% of the potential spermatozoa die spontaneously in the testis of adult mammals by the activation of an apoptotic programme (Blanco-Rodriguez, 1998). Although the cause(s) and the molecular mechanism(s) involved in germ cell apoptosis are far from defined, programmed cell death appears to be important to adjust germ cell numbers to that of the supporting Sertoli cells, and to ensure a quality control of the gametes produced (Braun, 1998). As judged by studies based on cell morphology and localization within the mammalian seminiferous tubule, germ cell degeneration occurs at any differentiative step involving spermatogonia, primary spermatocytes or spermatids (Oakberg, 1956; Clermont, 1962; Russell and Clermont, 1977; Huckins, 1978; Johnson et al., 1984; Kerr, 1992). When inter-nucleosomal cleavage of DNA—a characteristic feature of apoptosis (Wyllie, 1980)—is detected by in-situ end-labelling methods, apoptotic degeneration is observed in some primary spermatocytes and a few spermatogonia of adult normal rats (Billig et al., 1995; Blanco-Rodriguez and Martinez-Garcia, 1996; Brinkworth et al., 1998). DNA strand breaks have been identified in all types of germ cells in normal human testis (Hikim et al., 1998; Oldereid et al., 2001), although primary spermatocytes are the cells most commonly involved. These findings suggest that, in physiological conditions, the first meiotic division is the most critical step for germ cell apoptotic degeneration.
A quality control system, acting through apoptosis, has been suggested to be essential in mouse spermatocytes to monitor chromosome synapsis during meiosis. Indeed, an incomplete synapsis in pachytene cells is associated with a block in meiotic metaphase and subsequent germ cell loss through apoptosis, although the underlying molecular mechanism(s) has not been defined (Odorisio et al., 1998). Germ cell apoptosis may be induced by a variety of conditions, including gonadotrophin or growth factor withdrawal, irradiation, exposure to toxic or chemotherapeutic compounds, and cryptorchidism (Yoshinaga et al., 1991; Meistrich, 1993; Troiano et al., 1994; Billig et al., 1995; Henriksén et al., 1995, 1996; Hikim et al., 1995; Blanchard et al., 1996; Ling-Hong et al., 1996; Shetty et al., 1996; Blanco-Rodriguez and Martinez-Garcia, 1998; Yan et al., 2000). It is not known whether the same cellular signalling pathway activates germ cell apoptosis observed in different conditions. The Fas/Fas ligand (FasL) system seems to be implicated in apoptosis of germ cells. FasL is a type II membrane bound protein, which belongs to the tumour necrosis factor (TNF) family and is capable of inducing apoptosis in Fas-bearing cells (Suda et al., 1993; Tanaka et al., 1998). Within the mammalian testis, FasL expressed by Sertoli cells (Suda et al., 1993; French et al., 1996) induces apoptosis of Fas-expressing germ cells occurring either under physiological conditions or following different testicular injuries, such as radiation exposure or administration of Sertoli cells toxicants (Lee et al., 1997, 1999; Richburg et al., 2000). We have recently shown that Fas is expressed at both mRNA and protein levels by scattered germ cells in the adult human testis, while FasL is highly expressed by Sertoli cells, suggesting that, also in man, the Fas–FasL interaction might be involved in paracrine signalling between Sertoli cells and germ cells (Francavilla et al., 2000). In the present study, we have characterized the degenerative changes in germ cells by means of light (LM) and transmission electron (TEM) microscopy and correlated the frequency of degenerating germ cells with that of Fas-expressing germ cells in adult testes with normal, meiotic or post-meiotic arrest of spermatogenesis.
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Materials and methods |
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Testicular specimens
Archived testicular specimens were obtained from men with azoospermia or cryptozoospermia by open testicular biopsy. Informed consent was obtained from all patients. Each biopsy was divided into two parts: one was fixed in Bouin's solution, embedded in paraffin wax and used for immunohistochemistry, while the second part was immediately immersed in cold 0.1 mol/l cacodylate buffer, pH 7.4, containing 3% (v/v) glutaraldehyde for 2 h at 4°C. Samples were subsequently washed in cacodylate buffer and post-fixed in 1% (w/v) osmium tetroxide in distilled water, dehydrated through graded ethanol and embedded in Epon 812 (AGAR Scientific Ltd, Stansted, Essex, UK).
On the basis of standard qualitative interpretations of haematoxylin and eosin-stained paraffin sections, biopsies were classified as follows. (i) Normal histology: almost all tubules showing >10 elongating spermatids in each cross tubule section; (ii) Incomplete late maturation arrest (LMA): tubules showing spermatogenesis progressing through to elongated spermatids; the latter, however, were greatly reduced to less than five in each cross section of seminiferous tubule as well as tubules where only round spermatids are observed. (iii) Complete early maturation arrest of spermatogenesis (EMA): all tubules showing arrested spermatogenesis at the level of prophase of first meiotic division, with spermatids never observed.
Immunohistochemistry
Paraffin tissue sections of 5 µm were deparaffinized and subjected to immunohistochemical labelling as previously reported (Francavilla et al., 2000). Briefly, sections were incubated overnight at 4°C in phosphate-buffered saline containing polyclonal antibodies to Fas (N-18) at a concentration of 1 µg/ml (Santa Cruz Biotechnology, Santa Cruz, CA, USA). The Fas (N-18) antibody is an affinity-purified rabbit immunoglobulin (Ig)G raised against a peptide corresponding to amino acids 21–38 mapping at the amino terminus of the human Fas precursor. The antibody reacts with Fas of human origin by Western blotting and immunohistochemistry; it does not cross-react with other TNF superfamily type I transmembrane receptors, according to the supplier. Positive controls were obtained by testing the antibody on archived human hyperplastic lymphoid tissue (not shown) (Leithowser et al., 1993). The immunoreaction was detected by an avidin–biotin peroxidase complex method using a biotinylated goat anti-rabbit IgG (ImmunoPure ABC Peroxidase Rabbit IgG Staining Kit; Pierce, Rockfort, IL, USA). Sections were mounted with a mounting medium (Histovitrex; Carlo Erba, Milano, Italy) and visualized in a Leitz photomicroscope (Leica, Milano, Italy). Controls were performed by omission of the primary antibody or by incubating slides with the first antibody in the presence of inhibitor peptide (2 µg/ml) (Santa Cruz Biotechnology). Fas-positive germ cells were counted with the use of a x63 oil immersion objective lens and a x12.5 eyepiece, as reported (Francavilla et al., 2000). Longitudinal and cross sections of tubules with clear lumen were used for scoring. The total number of Fas-positive germ cells was divided by the total number of Sertoli cell nuclei, identified by counterstaining sections with haematoxylin, and the rate of positive cells was expressed as the number of Fas-positive germ cells per 100 Sertoli cell nuclei (FAS+GC%SE). Two-hundred Sertoli cell nuclei were counted in each biopsy.
Histology
Epon-embedded sections of 1 µm were stained in 1% (w/v) Toluidine Blue and analysed by light microscopy with a x63 oil immersion objective lens and a x12.5 eyepiece. The counting method was the same as that used for immunohistochemistry. Approximately 30 tubules were scored in each biopsy by counting, in each tubule, the number of elongating spermatids (SD), the number of degenerated germ cells (DG) and the number of Sertoli cell nuclei (SE). The total number of SD or DG was divided by the number of Sertoli cell nuclei (SD%SE or DG%SE).
Ultrastructure
Ultrathin 800–1200 Å sections were obtained from selected resin-embedded tissue blocks which, by LM, showed degenerating cells. Ultrathin sections were contrasted with uranyl acetate and lead hydroxyde (AGAR Scientific Ltd) and evaluated using a Philips M100 TEM (Philips Electronics, Eindhoven, Holland).
Statistical analysis
Kruskall–Wallis' one-way analysis of variance (ANOVA) by ranks was used to compare the number of FAS+GC%SE, and the number of DG%SE, in the three groups of biopsies with normal spermatogenesis (n = 10), LMA (n = 10) and EMA (n = 10). The small sample size suggested using non-parametric tests for data analysis. Post hoc comparison between pairs of groups was assessed by the Wilcoxon rank sum test, with a downward adjustment of the 
level to compensate for multiple comparisons. To maintain the overall probability at a level of 0.05 in the three independent comparisons, the value was divided by three to obtain a comparisonwise = 0.017 (0.05/3). Thus, each comparison was significant at the 0.017 level. The Wilcoxon rank sum test was also used to compare the number of SD%SE in the groups with normal spermatogenesis and with LMA. Correlations were evaluated with Spearman's rank correlation test. Data analysis was performed with SAS, version 6.12, 1996 (SAS Institute Inc., Cary, NC, USA).
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Results |
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Immunohistochemistry, histology and ultrastructure
In normal testes, Fas immunoreactivity was observed in Leydig interstitial cells, as already reported (Francavilla et al., 2000
), it was occasionally detected in the cytoplasm of isolated germ cells localized in the vicinity of the basal lamina (Figure 1A), and more often in the adluminal compartment of the seminiferous epithelium (Figure 1B). Immunostaining was observed in the cytoplasm of abnormal spermatids and in remnants of degenerating germ cells (Figure 1C), but was not detected in control sections in which the primary antibody was omitted (Figure 1D). On epon-embedded semithin sections, degenerating germ cells were identified as deeply stained pyknotic bodies (Figure 2A–C). An increased cytoplasmic basophilia, resulting in deep Toluidine Blue staining, might be an early change of apoptotic degeneration (Figure 2A) (Blanco-Rodriguez and Martinez-Garcia, 1998). Later apoptotic changes consisted of a diffuse cytoplasmic vacuolization associated with cell shrinkage, resulting in the formation of an empty pericellular space (Figure 2B). A deep homogeneous condensation of chromatin was observed at this stage, while more advanced apoptotic changes included a disassembly of the condensed nucleus into irregular masses of chromatin, dispersed in a huge pericellular space (Figure 2C). Degenerating cells could probably be identified as primary spermatocytes and round spermatids, since these changes were mostly localized in the adluminal compartment. Ultrastructural analysis confirmed that degenerative changes were observed in primary spermatocytes and in round or early elongating spermatids. Rare apoptotic spermatogonia were observed only in cases of deranged spermatogenesis and, as already reported, they were dislocated away from the basal lamina (Figure 3A) (Holstein et al., 1984).
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The identification as type A spermatogonia was indicated by the intermitochondrial cementing substance associated with the total absence of meiotic nuclear changes. Apoptotic type A spermatogonia showed chromatin condensation in the nuclear periphery with remaining large empty nuclear areas. The cytoplasm, containing mitochondria with dilated or disrupted cristae, was shrunken and an empty pericellular cleft indicated the partial/total loss of contact with Sertoli cells. Diplotene and pachytene spermatocytes were the germ cells that more frequently showed apoptotic features. Most degenerating cells were single cells, although occasionally a cluster of apoptotic cells was observed (Figure 3B
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In the early stage of apoptosis, meiotic nuclei displayed an exaggerated condensation of single chromosomes, showing an inner empty core containing remnants of synaptinemal complexes, formed by a single, thick, electron-dense fibril (Figure 3C,D). At this stage the nuclear membrane was only distorted or partially disrupted (Figure 3D) and the cytoplasm contained degenerating mitochondria and dilated endoplasmic reticulum (Figure 3C). In more advanced stages of apoptosis in meiotic cells, single condensed chromosomes were progressively replaced by an irregular network of condensed chromatin, while remnants of synaptinemal complexes progressively disappeared (Figure 3B). At this stage the nuclear and plasma membrane were grossly distorted and fragmented.
Early features of degeneration in spermatids were either restricted to the cytoplasm (Figure 4A), or involved both the cytoplasm and the nucleus, showing condensed chromatin in the perinuclear region (Figure 4B,C). These changes involved spermatids during Golgi and cup phases. In more advanced stages of spermiogenesis, condensed chromatin assumed a ring-like organization with a large intranuclear space containing fragmented or uncoalesed chromatin granules (Figure 4D–F). The destiny of these spermatids was either elimination in the seminiferous lumen (Figure 4D), or disposal through phagocytosis by Sertoli cells (Figure 4E,F). The latter was, however, quite rarely observed.
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Quantitative analysis of histology and immunohistochemistry
Table I
reports the numbers of degenerating germ cells and elongating spermatids referred to 100 Sertoli cell nuclei, evaluated on Toluidine Blue stained semithin sections, and the number of Fas-positive cells in testes with normal or deranged spermatogenesis. A significantly increased number of degenerating cells was observed in cases of incomplete LMA compared with normal testes and also to testes with complete EMA (P < 0.001 Wilcoxon's rank sum test). Testes with LMA also showed a greatly reduced number of elongating spermatids compared with normal testes (P < 0.001). A significantly increased number of Fas-positive germ cells was also observed in cases of LMA compared with normal testes (P < 0.001) and testes with EMA (P = 0.002). Complete early maturation arrest was associated with a huge variation in the rate of degenerating germ cells (range: 0.5–4.5/100 Sertoli cells) and of Fas-positive germ cells (range: 0.4–4.2/100 Sertoli cells). The number of degenerating germ cells/100 Sertoli cells was positively correlated with the number of Fas-positive germ cells/100 Sertoli cells (Figure 5A), while a negative correlation was found between the numbers of degenerating germ cells/100 Sertoli cells and of elongating spermatids/100 Sertoli cells (Figure 5B).
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Discussion |
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This study investigated the ultrastructure of degenerating germ cells in human testes with normal or deranged spermatogenesis, and correlated the number of degenerating cells with that of Fas-expressing germ cells. Ultrastructural analysis showed that apoptotic degeneration involved mostly primary spermatocytes and round or early elongating spermatids, while apoptotic spermatogonia were rarely observed and only in testes with abnormal spermatogenesis.
The identification of degenerating germ cells relies on nuclear and cytoplasmic changes as evidenced by LM, or on detection, in histological specimens, of internucleosomal cleavage of DNA by in-situ end-labelling methods (see Introduction). In both approaches, the identification of germ cell type undergoing degeneration is based mostly on cell morphology and its localization in the basal or in the adluminal compartment of the seminiferous tubule. By these criteria, spermatogonia, spermatocytes and round spermatids seem to be eliminated by apoptosis. Here, we show that the identification of degenerating germ cells in histological sections might be misleading. Apoptosis includes chromatin changes and cell shrinkage, which may hinder a correct identification of the step of spermatogenesis at which degeneration is triggered, due to the loss of cytological details and altered cellular size. Moreover, the ultrastructural identification of apoptotic primary spermatocytes in close vicinity to the basal lamina, and of apoptotic spermatogonia far from the basement membrane of the seminiferous epithelium, further questions the correct identification by LM, of the germ cell type involved in degeneration. On the whole, ultrastructural findings and analysis of epon-embedded sections at light microscopy suggest that apoptotic degeneration is an occasional finding in testes with normal spermatogenesis, as also shown by in-situ detection of DNA strand breaks (Oldereid et al., 2001). Apoptosis seems to involve mostly primary spermatocytes and spermatids during early differentiation and only rarely spermatogonia. This degenerative programme was activated in a larger number of germ cells in cases of an incomplete post-meiotic arrest of germ cell maturation, or in selected cases of complete meiotic arrest of spermatogenesis. The increased number of apoptotic cells in cases of incomplete post-meiotic arrest of spermatogenesis and its inverse correlation with the number of elongating spermatids, suggests that a programmed death of germ cells might contribute to the low efficiency of spermatogenesis. However, death activation of primary spermatocytes and round spermatids is not the only defect observed in cases of incomplete post-meiotic arrest of spermatogenesis. A ubiquitous incomplete chromatin condensation of the few spermatids which reach a full elongation in stage I of the seminiferous epithelium has been shown in this condition (Francavilla et al., 2001). Normal chromatin condensation during spermiogenesis (Hecht, 2000) as well as apoptosis of germ cells (Blanco-Rodriguez, 2000) result from a regulated, ordered activation of gene expression, suggesting that specific patterns of altered expression of testis-specific genes might be responsible for both impaired differentiation and activated apoptosis of germ cells.
Inactivation of the cAMP-responsive element modulator (CREM) gene—one of the genes involved in spermatid differentiation (Tamai et al., 1997)—is associated with a post-meiotic arrest of maturation and with a several-fold increase in the number of apoptotic germ cells (Nantel et al., 1996). The same histological pattern has been reported in human testes which do not express the activation isoform of the CREM transcript (Peri et al., 1998; Weinbauer et al., 1998; Steger et al., 1999). Animal models have demonstrated a total block of the first meiotic division and an elimination by apoptosis in spermatocytes with synaptic errors (Odorisio et al., 1998). The atypical synaptonemal complexes observed in apoptotic pachytene spermatocytes (Figure 3C,D) might be indicative of an altered homologous chromosome pairing and consequent activation of apoptotic disposal of affected cells. A total block of the first meiotic division and associated apoptotic activation of spermatocytes has been observed in mice deficient for numerous gene products implicated in homologous recombination at meiosis (Venables and Cooke, 2000). All these data demonstrate that different conditions of altered spermatogenesis resulting in a complete block of maturation during the first meiotic division, or in an incomplete post-meiotic arrest of maturation, are associated with an activation of the apoptotic disposal of primary spermatocytes and spermatids.
The tight correlation observed between the number of apoptotic cells and the number of Fas-expressing germ cells in all testicular specimens, suggests that an altered execution of the genetic programme required for a regular meiotic and post-meiotic germ cell maturation might be associated with an up-regulation of Fas gene expression in germ cells. The cross-linking of the Fas receptor with the Fas-ligand expressed by Sertoli cells (Francavilla et al., 2000) might trigger the apoptotic degeneration of defective human germ cells. The molecular control of Fas gene expression in such different conditions of altered germ cell maturation spanning from complete meiotic arrest to an incomplete post-meiotic arrest of spermatogenesis is totally unknown. The few data available suggest that Fas gene expression in lymphocytes during Fas-induced cell death seems to be regulated by transcriptional activators and repressors (Rudert et al., 1998; Lasham et al., 2000), or by protein kinase C, through a regulatory gene TDAG51 (Wang et al., 1998). Future studies should define whether or not these molecular mechanisms of Fas gene control are operating in normal germ cells as well as in germ cells that harbour a defective differentiation programme.
The molecular mechanisms which mediate the clearance of apoptotic cells are numerous and not yet well-characterized (Savill and Fadok, 2000). In-vitro studies have shown that rat Sertoli cells phagocytose spermatogenic cells by recognizing phosphatidylserine exposed on the surface of apoptotic cells through a class B scavenger receptor type 1 (Shiratsuchi et al., 1999). Ultrastructural findings reported here support an uncommon occurrence of Sertoli cell phagocytosis of apoptotic germ cells, as well as a more frequent precocious spermiation of degenerating spermatids in the lumen of the seminiferous tubule. Apoptotic degeneration in tissue sections was a scattered phenomenon in cases of deranged spermatogenesis and this might hamper easy demonstration and identification of apoptotic cells eliminated by phagocytosis in human testis. On the other hand, it has been claimed that apoptotic spermatocytes and spermatids might not be easily removed by phagocytosis in the human testis with spermiogenic failure (Tesarik et al., 1998). It remains to be defined whether and which molecular mechanism is activated in vivo in the human testis, to mediate the apparently rapid and efficient elimination of apoptotic germ cells observed in the rat, by in-vitro models. This might have some relevance in the era of testicular extraction of spermatids and subsequent ICSI to treat infertility for azoospermic men. The increased activation of spermatid apoptosis, not followed by a rapid clearance, might contribute to an impaired potential for embryo development, after ooplasmic injection of testicular immature (Tesarik et al., 1996; Sofikitis et al., 1998) or elongated spermatids (Francavilla et al., 2001) reported in cases of non-obstructive azoospermia.
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We are grateful to Teresa Odorisio for her critical reading of the manuscript. This work was supported by MURST, Italy.
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4 To whom correspondence should be addressed at: Department of Internal Medicine, Andrology Unit, University of L'Aquila,Via S. Sisto 22E, 67100 L'Aquila, Italy. E-mail: Sandrof{at}univaq.it
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Submitted on July 17, 2001; accepted on December 6, 2001.
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