Involvement of Germ Cell Apoptosis in the Induction of Testicular Toxicity Following Hydroxyurea Treatment

Toxicology and Applied Pharmacology 155, 139 –149 (1999) Article ID taap.1998.8593, available online at http://www.idealibrary.com on

Involvement of Germ Cell Apoptosis in the Induction of Testicular Toxicity Following Hydroxyurea Treatment Jae-Ho Shin,*,† Chisato Mori,* and Kohei Shiota*,‡ *Department of Anatomy and Developmental Biology and ‡Congenital Anomaly Research Center, Faculty of Medicine, Kyoto University, Kyoto, Japan; and †Toxicology Research Institute, Korea Food and Drug Administration, Seoul, Korea Received August 4, 1998; accepted November 5, 1998

Involvement of Germ Cell Apoptosis in the Induction of Testicular Toxicity Following Hydroxyurea Treatment. Shin, J.-H., Mori, C., and Shiota, K. (1999). Toxicol. Appl. Pharmacol. 155, 139 –149. The present study investigated the occurrence of apoptotic cell death in the mouse testis at various intervals following the administration of hydroxyurea (HU). The presence of apoptosis was assessed by the terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end labeling (TUNEL) method and by DNA fragmentation assay using ligation-mediated polymerase chain reaction. Both the incidence of apoptotic cells and the level of DNA fragmentation in the testis increased depending on the HU dose, and they were most apparent at the highest dose (400 mg/kg). The incidence of apoptotic cells in the HU-treated group increased continuously and peaked at 12 h, but then decreased gradually, reaching control levels by 48 h. After HU treatment, TUNEL-positive apoptotic cells increased in the seminiferous epithelium of the tubules, and affected cells were found synchronously in the tubules of animals treated with HU. Spermatogonia and spermatocytes were found to be affected selectively. TUNELpositive cells were found to be stage-specific and were primarily in stage IV–VI tubules. It has been shown that in vivo HU exposure induced testicular germ cell apoptosis dose dependently in a timeand stage-specific manner, and damaged cells appeared to be eliminated by phagocytosis by neighboring cells. Apoptosis of damaged testicular germ cells is apparently a common response to various testicular toxicants therefore protecting the next generations of germ cells from the damaged cell population. © 1999 Academic Press

Key Words: hydroxyurea; germ cell apoptosis; mouse testis.

In mammalian spermatogenesis, three principal phases of cellular differentiation can be discerned. During the initial phase, spermatogonia proliferate and differentiate to give rise to spermatocytes in the basal compartment of the seminiferous epithelium. During the second phase, spermatocytes form haploid spermatids by means of two consecutive meiotic divisions, while primary and secondary spermatocytes remain intercalated by the cytoplasmic processes with adjacent Sertoli cells. The final phase of spermatogenesis, which is referred to as

spermiogenesis, is the process of morphological transformation of haploid cells (the spermatids) to spermatozoa. In the mouse, seminiferous tubules are classified into 12 stages according to the morphologically specific features of differentiating germ cells of the seminiferous epithelium (Oakberg, 1956; Russell et al., 1990). As in many other tissues, the size of the spermatogenetic cell population in the testis appears to be controlled not only by cell proliferation but also by a dynamic balance with cell death (Huckins, 1978). Germ cell degeneration in the mature testis occurs spontaneously during normal spermatogenesis, and a loss of early germ cells by apoptotic degeneration is considered to contribute to the control of the size of the spermatid population (Huckins, 1978; Kerr, 1992). On the other hand, germ cell apoptosis can also be induced by pathological conditions such as hypophysectomy (Tapanainen et al., 1993), withdrawal of testosterone by Leydig cell depletion (Troiano et al., 1994), cryptorchidism (Shikone et al., 1994), and exposure to X irradiation (Henriksen et al., 1996; Hasegawa et al., 1997), heat (Shikone et al., 1994) or various chemical agents (Blanchard et al., 1996; Cai et al., 1997; Ku et al., 1995; Nakagawa et al., 1997; Richburg and Boekelheide, 1996; Strandgaard and Miller, 1998). Apoptosis is a morphologically distinct form of programmed cell death that plays a major role in development, homeostasis, and various diseases (Steller, 1995). Apoptotic cell death occurs through the activation of a cell-intrinsic suicide program to remove potentially damaged cells, and the activation of the program is regulated by many different signals that originate from both the intracellular and the extracellular milieu. Such evolutionary conserved cell death through apoptosis is distinguished from pathological cell death or necrosis by their characteristic morphological and biochemical features (Kerr et al., 1972; Wyllie, 1987). By activation of endogenous endonucleases which cleave the genomic DNA periodically, nuclear DNA of apoptotic cells is degraded into fragments which can be detected as a characteristic DNA ladder on gel electrophoresis (Wyllie, 1980). Hydroxyurea (HU) exhibits cytotoxic effects on the rapidly proliferating S-phase cells and ultimately induces cell death by

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specific inhibition of DNA synthesis without any effect on RNA or protein synthesis. In Xenopus embryos, treatment with HU arrests the cell cycle at interphase and induces widespread apoptosis that occurs coincidentally with cyclin A degradation at the equivalent of the early gastrulation transition (Stack and Newport, 1997). Yabro (1992) showed that HU exerts its anti-proliferative effects by inactivating ribonucleotide reductase which is a key enzyme for the transformation of ribonucleotides to the deoxyribonucleotides essential for DNA synthesis. Because of its cytotoxic effect, HU has been used alone or with other chemicals or X-ray for chemotherapeutic treatment for chronic myelocytic leukemia and other hematological malignancies (Kennedy, 1972) and meningiomas (Schrell et al., 1997). It is reported recently that HU therapy can ameliorate the clinical course of sickle cell anemia (Charache et al., 1995) and is also applied to children and young adults (Ferster et al., 1996; Shetty et al., 1998). In addition, HU is noted to exhibit potent synergism by its ribonucleotide reductase inhibitory activity with the anti-HIV-1 dideoxynucleoside 29,39-dideoxyinosine, bringing about failure of HIV DNA synthesis and, thus, of HIV replication (Gao et al., 1998). When given as single injections, HU is cytotoxic to spermatogenic cells and exerts harmful effects on reproduction (Lu and Meistich, 1979). HU exposure alters mouse testicular cell kinetics and sperm chromatin structure when injected for 5 days (Evenson and Jost, 1993). Wiger et al. (1995) showed that intraperitoneal injections with HU (200 mg/kg) to male mice for 5 consecutive days induced atrophy of seminiferous tubules 5 days after the last injection. Flow cytometry revealed marked reduction in the tetraploid population of testicular cells, mostly in early pachytene spermatocytes, followed by later changes in the population of spermatids at various stages of maturation. However, it remains to be elucidated whether apoptotic cell death is involved in HU-induced testicular toxicity. The present study was undertaken to examine the occurrence of apoptosis in male germ cells in mice following HU treatment. We examined the dose dependence of apoptotic cell death and its sequential changes in the testis by biochemical DNA fragmentation assay for DNA ladders and by in situ cytochemical detection of apoptotic cells by the terminal deoxynucleotidyl transferase (TdT)-mediated dUTP-biotin nick end labeling, the TUNEL method. The correlation between the occurrence of apoptosis and the stage of germ cell differentiation was also analyzed. MATERIALS AND METHODS Treatment of animals. Male ICR mice were purchased from SLC (Shizuoka, Japan) and housed in a temperature and humidity controlled room with free access to food and water on a 12-h light/dark cycle (darkness from 8 p.m. to 8 a.m.). The mice used in this study were 6 –7 weeks old and 30 –37 g at the start of the experiment. They were divided randomly into groups, weighed, and injected ip with HU (Wako Pure Chemical, Osaka, Japan), and control animals were injected with physiological saline (vehicle) according to the experimental schedule. To avoid general toxic effects caused by higher doses, a dose of 400

mg/kg HU was the maximum used in the time course study. The animals were killed and their testes examined at 0, 4, 8, 12, 18, 24, and 48 h after HU treatment. A dose-response study was carried out at 12 h after treatment, using 0, 100, 200, and 400 mg/kg HU. After weighing both testes of each animal, the left testis was fixed in 4% phosphate-buffered paraformaldehyde solution (pH 7.4) for histological studies, and the right was frozen in liquid nitrogen and stored at 280°C until DNA fragmentation assay was performed. Terminal deoxynucleotidyl transferase (TdT)-mediated dUTP-biotin nick end labeling (TUNEL) assay. The fixed testes were dehydrated through a graded series of ethanol and embedded in paraffin according to standard procedures. Paraffin sections (5 mm thick) were placed on slides pretreated with 3-aminopropyltriethoxysilane and stored at room temperature until further processing. The in situ visualization technique (TUNEL) was carried out according to the method of Mori et al. (1997) which was a modification of that of Gavrieli et al. (1992). After deparaffinization and rehydration, the sections were treated with 20 mg/ml proteinase K (Wako Pure Chemical, Osaka, Japan) at 37°C for 10 min. After proteinase K treatment, the labeling was carried out as previously reported (Mori et al., 1997). The sections were covered with the Vectastain Standard ABC reagent (Vector, Burlingame, CA) for 30 min at room temperature, washed in PBS, and then developed with 3,39 diaminobenzidine (DAB). The slides were counterstained, dehydrated through a graded series of ethanols, and coverslipped for observation under a microscope. For each testis, cross sections of at least 100 tubules were scored using routine light microscopy. To determine the stage specificity of TUNEL staining, we performed periodic acid Schiff (PAS) staining and classified the seminiferous tubules into four groups, i.e. stages I–III, IV–VI, VII–IX, and X–XII according to the staging criteria described by Oakberg (1956) and Russell et al. (1990). TUNEL-positive germ cells were identified as spermatogonia, spermatocytes, or spermatids by their location in the seminiferous epithelium, PAS-stained acrosome, and the features of neighboring cells. When applicable, the number of apoptotic germ cells per tubule was counted for each stage group. To identify the cell type associated with TUNEL-positive staining in the tubules, a double immunohistochemical staining was undertaken using the anti-mouse heat shock protein 70-2 polyclonal antibody (HSP70-2 [a rabbit IgG] provided by Dr. E. Eddy, NIEHS, USA) which is a marker for late spermatocytes and spermatids (Rosario et al., 1992). To detect the marker protein and DNA fragmentation in TUNEL-positive cells, FITC-conjugate goat anti rabbit IgG (TAGO Inc., CA) was used as a secondary antibody for the HSP70-2, and rhodamine B-labeled streptavidin (Molecular Probes Inc., OR) for the biotin-labeled DNA fragmentation (Mori et al., 1997). DNA fragmentation assay by ligation-mediated polymerase chain reaction (LM-PCR). For the DNA fragmentation assay, low molecular weight DNA was isolated by the method adapted from Tilly and Hsueh (1993). After thawing the testes, homogenization buffer (0.1 M NaCl, 10 mM NaEDTA [pH 8.0], 0.1 M Tris HCl [pH 8.0], 0.2 M sucrose) was added. The testes were homogenized with 50 strokes in a homogenizer. To each homogenate, 12 ml of 10% SDS was added and then incubated for 30 min at 65°C. Following the addition of 35 ml of 8 M potassium acetate to each sample to precipitate the proteins, the samples were placed on ice for 60 min and then centrifuged. The collected supernatants were then extracted with phenol:chloroform:isoamyalcohol (25:24:1) and then chloroform:isoamyalcohol (24:1), precipitated with ethanol, and resuspended. RNA in each sample was digested with 1 ml (0.5 mg/ml) DNase-free RNase for 60 min at 60°C (10 mM Tris, pH 7.5, 15 mM NaCl). The samples were again extracted, precipitated with ethanol, and dried. To visualize nucleosomal ladders for fragmented DNA and to compare the amount of DNA fragmentation between groups, LM-PCR modified from Staley et al. (1997) was undertaken. DNA ladders were detected using the ApoAlert LM-PCR Ladder Assay Kit (CLONTECH, CA) according to the manufacturer’s protocol. Briefly, 2 mg of extracted DNA from each testis was incubated to anneal and ligate with the adaptor oligonucleotides. The adaptorligated DNA was used for PCR. PCR products (10 ml of each reaction) were electrophoresed on a 1.2% agarose gel for 30 min at 50 V. DNA was stained with ethidium bromide and visualized with an ultraviolet transilluminator. Resulting ladders should comprise bands at intervals of approximately 200 bp

HYDROXYUREA-INDUCED APOPTOSIS IN MOUSE TESTIS because of the addition of the adaptors (48 bp). The size of the resulting DNA bands was estimated on the base of a 100-bp DNA ladder standard marker (Pharmacia, NJ). A computer-connected, scanned image of each film was created, and then analyses were performed using the public domain NIH Image program (developed at the U.S. National Institutes of Health and available on the Internet at http://rsb.info.nih.gov/nih-image/). For each lane the total integrated optical density (IOD) of the lane was used as a measure of low molecular weight DNA to compare the relative amounts of DNA fragmentation in each sample. Statistical evaluation. The significant correlation between the groups was determined by one-way analysis of variance (ANOVA). Statistical analysis was undertaken using a computer software program (StatView 4.01). The post hoc comparisons were made using the Scheffe test. The difference between groups was considered significant at the level of p , 0.05. All the results are presented as mean 6 SE.

RESULTS

Body and Testis Weights At any of the time intervals during the experimental period, no significant differences in body and testis weights between the control and HU-treated groups were found. Further, no dead animals were recorded in any group. Also the animals treated with HU did not show any specific toxic signs related to the treatment. Morphologic Characteristics of Apoptosis in Testis Germs after HU Treatment Figure 1 shows apoptotic cells in the mouse testis as assessed by the TUNEL method. Apoptotic cells in the seminiferous tubules was increased in HU-treated animals when compared with controls (Figs. 1A and 1B). Apoptotic cells identified by TUNEL-positive nuclei were found in the peripheral region near the basement membrane of seminiferous tubules. In both the control and HU-treated animals, most apoptotic cells were confined to the spermatogonium and spermatocyte populations. There were no TUNEL-positive cells in the interstitial tissue in both controls and HU-treated mice. Compared with the control mouse testis (Fig. 1A), considerably more apoptotic cells were found synchronously in damaged tubules of HU-treated animals (Fig. 1B). Most of the TUNEL-positive cells were observed in the areas where the HSP70-2 was not expressed (Fig. 1C). The frequency of TUNEL-positive cells was variable among the tubules of HUtreated mice (Fig. 2). However, multinucleated giant cells in damaged tubules did not show TUNEL-positive staining. Except for the increase in apoptotic cells, the histological architecture of the seminiferous tubules appeared normally maintained after HU treatment. Gel Fractionation of DNA from Testis after HU Treatment To examine whether DNA ladders were observed in the testis of animals treated with HU, we examined the DNA fragmentation patterns in each group by LM-PCR. As illustrated in Fig. 3A, the fragmented DNA was observed as a

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ladder-like appearance in both the control and HU-treated animals. The intensity of DNA ladders increased with the HU dose: the ladder was most clearly recognized at the highest dose, 400 mg/kg. In visual quantitation using the NIH image program, no significant differences were noted in the IOD of DNA ladders between the control group and groups treated with 100 or 200 mg/kg HU (Fig. 3B). The intensity of DNA ladders significantly increased in the 400 mg/kg group when compared with the controls. In addition, the degree of DNA fragmentation was significantly increased, by 400 mg/kg HU when compared with the 100 or 200 mg/kg HU groups. To compare the histologically detected cell death with the onset of DNA cleavage for apoptosis, a time-course analysis of HU-induced DNA fragmentation was undertaken. Figure 4A demonstrates the PCR products of isolated DNA at various time intervals after HU treatment. Although DNA ladders were detected all time points, they were most pronounced at 12 h after HU treatment. However, DNA ladders at 18, 24, and 48 h became less intense than those at 12 h following HU treatment. A quantitative analysis showed that the IOD of DNA ladders was significantly increased in the testes at 12 h after exposure to 400 mg/kg of HU. The IOD of DNA ladders decreased significantly thereafter, and returned to the control level by 48 h (Fig. 4B). Assessment of in Situ DNA Fragmentation in Testis Germ Cells after HU Treatment In the dose-response experiment, animals were treated with doses of 0, 100, 200, or 400 mg/kg of HU and killed 12 h later because TUNEL-positive cells were observed most abundantly in the time course experiment described previously. The number of seminiferous tubules containing apoptotic germ cells significantly increased with the HU dose (Fig. 5A). The incidence of apoptotic cells increased dose dependently following HU treatment in all the tubules and in damaged tubules (Fig. 5B). Further the number of apoptotic germ cells in each group was calculated as an average for 100 tubules. There was no significant difference between the control and 100 mg/kg HUtreated groups in the mean number of apoptotic germ cells observed in the seminiferous epithelia of cross sections of all the tubules or the tubules containing at least one apoptotic cell. The highest dose of HU, 400 mg/kg, induced a significant increase in the incidence of apoptosis, when compared with the control and lower dose groups. Figure 6 shows a time course of the occurrence of apoptosis in response to a single dose of 400 mg/kg HU. The number of seminiferous tubules containing apoptotic germ cells increased with time until 12 h after treatment, and then decreased gradually to control levels by 48 h (Fig. 6A). Figure 6B shows the number of apoptotic cells in all tubules observed and shows that in the tubules containing any apoptotic cells, the relative incidence of apoptotic cells to the total germ cells in the tubules increased after HU treatment, peaking at 12 h at a level about

FIG. 1. Cross sections of the seminiferous tubules of the mouse testis. The tubules are double-stained with TUNEL and HSP70-2. Red fluorescent TUNEL-positive cells (arrowheads) are detected by rhodamine B and green fluorescent HSP70-2 in the luminal region are detected by FITC. Few TUNEL-positive apoptotic cells are found in the controls (A), but numbers increased in affected tubules 12 h after treatment of 400 mg/kg HU (B). TUNEL-positive cells are in the region near the basement membrane of the seminiferous tubules. Double staining of the same tubule by TUNEL and HSP70-2 demonstrates that most of the TUNEL-positive cells are observed in the area where HSP70-2 is not localized (C). Bar, 50 mm.

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FIG. 2. Apoptotic cells in the seminiferous tubules of the mouse testis 12 h after HU treatment. Testicular tubules of stages I–III, IV–VI, VII–IX, and X–XII, respectively, in HU-treated male mice (a– d). Tubules were staged by PAS-stained acrosomal vesicles (arrows) and mitotic figures (m), and apoptotic cells were detected by DAB (arrowheads). TUNEL-positive apoptotic cells, stained black, are predominantly spermatogonia and early spermatocytes at stages I–III, and IV-VI of the seminiferous tubules. Bar, 20 mm.

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FIG. 3. Dose response of HU-induced apoptosis by DNA fragmentation assay in the testis of control and HU-treated mice. (A) DNA ladders observed by LM-PCR. The molecular markers (100-bp ladder) are on lane M. Lanes 1– 4 represent LM-PCR products from extracted DNA from the mouse testis 12 h after treatment of 0 (vehicle only), 100, 200, and 400 mg/kg HU, respectively. (B) A histogram depicting the total fragmented DNA determined by image analysis of photographs such as that shown in 3A. Values are mean 6 SE of 3 separate experiments. *, significantly different from the control (0 mg/kg, p , 0.05); #, significantly different from the 100 mg/kg group ( p , 0.05); ˆ, significantly different from the 200 mg/kg group ( p , 0.05).

4-fold above the control, and then decreasing gradually to return to control levels by 48 h. There were no significant differences among 0-, 4-, 8-, 24-, and 48-h groups of HUtreated animals in the mean number of apoptotic germ cells per total tubules or the tubules containing any apoptotic cells. However, apoptotic germ cells increased significantly at 12 and 18 h after HU treatment, when compared with control and the other time points. In comparison with the 12-h data, signifi-

cantly fewer apoptotic germ cells were found in apoptotic tubules at 18, 24, and 48 h after HU treatment. A quantitative analysis was undertaken on the designated stage groups to examine the relationship between the spermatogenetic stage and the susceptibility to induced apoptosis (Fig. 7). For quantitative analysis the observed tubules were divided into 4 groups depending on the number of apoptotic cells per tubule, i.e., 0, #3, #7, and .7. For all stage groups, no

FIG. 4. Time course study by DNA fragmentation assay for the testis of control and HU-treated mice. (A) DNA ladders observed by LM-PCR. The molecular markers (100-bp ladder) are on lane M. Lanes 1–7 represent LM-PCR products from extracted DNA at 0, 4, 8, 12, 18, 24, and 48 h, respectively, following treatment with 400 mg/kg HU. (B) A histogram depicting the total fragmented DNA determined by image analysis of photographs such as that shown in 4A. Values are mean 6 SE of 3 separate experiments. *, significantly different from the control (0 h, p , 0.05); #, significantly different from 12 h after HU treatment ( p , 0.05).

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FIG. 5. Dose response for apoptosis in the mouse testis at 12 h after HU treatment. (A) The percentage of seminiferous tubules containing one or more TUNEL-positive cells. (B) The number of apoptotic germ cells in all the tubules observed (■) and within TUNEL-positive seminiferous tubules (h). Apoptotic cells are presented, respectively, as the number of apoptotic cells per 100 tubules in each group. Values are mean 6 SE. *, significantly different from the control (0 mg/kg, p , 0.05); #, significantly different from the 100 mg/kg group ( p , 0.05); ˆ, significantly different from the 200 mg/kg group ( p , 0.05).

significant statistical differences were observed in the proportion of apoptotic tubules per all tubules between the control (0 h) and the 4-h point. Although the difference between the control and HU-treated groups was not significant at any time point for tubules at stages VII–IX, an increase in the occurrence of affected tubules was observed in stages I–III, IV–VI, and X–XII, with a peak at 12 h. Specifically, tubules with more than 7 apoptotic cells were markedly increased in stages I–III and IV–VI at 12 h after HU treatment.

DISCUSSION

In this study, it has been demonstrated that HU can induce apoptosis in mouse male germ cells in a dose-dependent pattern, and this is the first reported demonstration that a single dose of HU induces apoptotic death of male mouse germ cells. A DNA fragmentation assay using LM-PCR also demonstrated that germ cell apoptosis induced by HU reaches a maximum peak at 12 h after treatment. Further HU increased the fre-

FIG. 6. Time course of the occurrence of apoptosis induced by HU (400 mg/kg) in the mouse testis. (A) The percentage of seminiferous tubules containing one or more TUNEL-positive cells. (B) The number of apoptotic germ cells in the all tubules observed (■) and within TUNEL-positive seminiferous tubules (h). Apoptotic cells are presented, respectively, as the number of apoptotic cells per 100 tubules in each group. Values are mean 6 SE. *, significantly different from the control (0 h, p , 0.05); #, significantly different from 12 h after HU treatment ( p , 0.05).

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FIG. 7. A comparison of the relative incidence of apoptotic tubules in various stages of the seminiferous epithelial cycle at different time intervals after treatment with 400 mg/kg HU. Apoptotic cells at each stage were recorded separately (n 5 3 for each group). Throughout the experimental period, there were no tubules which could not be staged. Data are presented as percentage of apoptotic tubules with apoptotic cells per tubule, respectively, #3 (■), #7 (p), and .7 (u).

quency of TUNEL-positive cells and also the number of the tubules containing affected germ cells. These changes returned to the control level by 48 h after treatment. On the gel electrophoresis for DNA ladders, a significant increase in DNA fragmentation was observed at 12 h after HU treatment when TUNEL-positive cells increased significantly. Since DNA fragmentation increased at 8 h after HU treatment, DNA fragmentation may be initiated as early as 8 h after exposure to HU. An increase in germ cell apoptosis in rodents has also been reported to occur in response to other testicular toxicants (Blanchard et al., 1996; Cai et al., 1997; Ku et al., 1995; Strandgaard and Miller, 1998) and to the withdrawal of testosterone (Billig et al., 1995; Sinha Hikim et al., 1995; Troiano et al., 1994), but in the tubules of control animals also a small number of spontaneous apoptotic cells has been observed as a part of the normal process of spermatogenesis (Brinkworth et al., 1995; Kerr, 1992; Nakagawa et al., 1997), a finding confirmed by this study. Spontaneous germ cell apoptosis is detected mainly in premeiotic cells (Billig et al., 1995; Cai et al., 1997). During spermatogenesis, germ cells that are either spontaneous or chemically initiated into a mutagenic or precarci-

nogenic state are known to be recognized and removed by apoptosis. We speculate that apoptosis of germ cells may be a mechanism by which the cells undergoing abnormal and irreciprocal changes are detected and destined to die. HU has been reported to induce cell death in vitro via inhibition of ribonucleotide reductase, producing a depletion of necessary DNA precursors (Skog et al., 1987). In this study, HU-induced apoptosis was found only in germ cells but not in interstitial cells or Sertoli cells. It is important to note that HU preferentially affects germ cells in damaged tubules in a synchronous fashion since the increase in apoptotic cells was more evident in affected tubules than in the total tubules. This corresponds to the results reported previously for apoptosis induced by various chemicals. For example, treatment with a GnRH antagonist, azaline-B, increased germ cell apoptosis in apoptotic tubules, and the apoptotic cells were identified as spermatocytes in the region near the basement membrane of the tubules (Billig et al., 1995). In cyclophosphamide-treated rats, a high incidence of apoptotic cells was found synchronously in the seminiferous tubules (Cai et al., 1997). These observations could be explained by the fact that spermatogonia and spermatocytes are connected by intercellular bridges which serve to transduce intracellular signals and contribute to synchronizing their differentiation (Clemont, 1972). It appears that HU-induced apoptosis of germ cells plays a protective role so that damaged germ cells may be recognized and efficiently removed by apoptosis during spermatogenesis. The mean number of apoptotic germ cells in all the observed tubules increased as a function of time after HU treatment and the number of apoptotic germ cells within apoptotic tubules also increased and peaked at 12 h after treatment. The time for apoptosis to peak in response to chemicals varies, depending on the cell type and the type of the stimulus (Stephens et al., 1993). For example, hyperthermia induces apoptosis in 30 min, methotrexate at 18 h, and 1,3-dinitrobenzene at 24 h in somatic cells after treatment (Barry et al., 1990). Cyclophosphamide induces maximal apoptosis in Oca-I tumor cells between 12 and 18 h, but in Mca-4 tumor cells at between 8 and 12 h after treatment (Meyn et al., 1994). An in vivo study on cyclophosphamide-induced apoptosis showed that the maximal induction of apoptosis was observed at 12 h in the tubules of the rat testis (Cai et al., 1997) which is similar to the results presented in this study. Pharmacokinetics, metabolic activation, and signaling pathways for apoptosis vary between chemicals and may explain the difference in the time course of the apoptotic response. In the present study, the number of apoptotic cells decreased gradually after the peak and returned to normal levels 48 h later. After attaining the maximum levels of apoptosis by the administration of 1,3-dinitrobenzene (Strandgaard and Miller, 1998) or cyclophosphamide (Cai et al., 1997), the incidence of apoptotic cells was reported to decrease rapidly to control levels. Apoptosis is a relatively rapid mechanism of cell death, and complete removal of the cell from the tissue often occurs within hours (Bursch et al., 1990; Jacobson et al.,

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1997). The incidence of apoptotic cells decreases at later time points probably because the neighboring Sertoli cells eliminate apoptotic germ cells by phagocytosis. Our dose-response study showed that DNA ladders increased with the dose of HU and were most obvious in the 400 mg/kg HU group, which was similar to the results of the TUNEL study. Since HU doses of 200 mg/kg or less did not induce significant amounts of apoptosis, these doses of HU appeared to be below the threshold level to induce lethal damage of germ cells. A 5-day consecutive treatment with 200 mg/kg HU produced testicular atrophy, and some tubules were virtually devoid of the seminiferous epithelium, containing mainly Sertoli cells 5 days after the last treatment (Wiger et al., 1995). In this study with a single dose of HU, however, there were no mice with reduced testicular weight or signs of irreversible tubular atrophy up to 14 days after a single dose of 400 mg/kg HU treatment. These observations suggest that multiple doses of HU, although lower than those used in this study, can induce continued depletion of spermatogenetic cells and subsequently result in reduced testicular weight and irreversible testicular atrophy. When damaged cells are eliminated by apoptosis after a single dose of HU, surviving germ cells may recover from damage and/or transient mitotic or meiotic arrest and restore spermatogenesis soon after treatment. The present study showed that germ cells at stages IV–VI appeared to be the most sensitive group to HU-induced apoptosis, and spermatogonia and spermatocytes were the primary cell populations undergoing apoptosis. To date there were several reports on the chemical-induced apoptosis of male germ cells, but only a few of them analyzed celland stage-specificity. In somatic cells, each cell line varies in the apoptotic response to cytotoxic stimuli, and its sensitivity may depend on the differences in the expression of genes related to apoptosis or cell differentiation. Germ cells at different stages of the seminiferous epithelium are known to possess their own way to differentiate and respond differently to the stimuli, although the mechanisms of chemically induced stage-specific apoptosis remains unclear. In the present study, no significant difference was observed between the control and HU-treated groups concerning apoptosis in stage VII–IX tubules, but affected tubules increased at stages I–III, IV–VI, and X–XII following HU treatment. Such stage-specific occurrence of apoptotic cells also varies among different chemicals. Treatment with gonadotropin antagonists, azaline-B (Billig et al., 1995), or ethane dimethane sulfonate (Henriksen et al., 1995; Troiano et al., 1994) increased apoptosis, predominantly in spermatocytes and spermatids at stages VII and VIII, but the cells recovered from the damage with testosterone supplementation. After 1,3-nitrobenzene treatment, extensive damage of seminiferous tubules was caused by degeneration of pachytene spermatocytes at stages VII through VII (Hess et al., 1988). Cyclophosphamide-induced apoptosis in rats was reported to be most pronounced in spermatogonia and sper-

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matocytes at stages I–IV and XI–XIV, but did not increase markedly at stages V–VII and VII–X (Cai et al., 1997). Testosterone is known to preferentially affect the cells at stages VII–VIII of the seminiferous epithelium and to have an important effect on the conversion of step 7– 8 spermatids (O’Donnell et al., 1994). Therefore, it is unlikely that hormonal changes can be a cause of the observed cell- and stage-specificity of apoptosis in testicular germ cells after treatment of HU. In this study, HU-induced apoptotic cells were markedly increased at stages IV–VI, when spontaneous or cyclophosphamide-induced apoptosis occurs infrequently. When the rate of testicular DNA synthesis was studied in the rat at various time intervals after a single dose of chemotherapeutic agents, HU, a non-DNA damaging agent, produced diverse patterns of DNA changes that are different from those caused by cyclophosphamide, which is a DNA damaging agent (Lambert and Eriksson, 1979). Thus, the cell- and stage-specific mode of apoptotic induction in the seminiferous epithelium differs by the chemical type used, probably due to the difference in its activity and mechanisms of action and in the susceptibility of male germ cells at different stages of spermatogenesis. This study demonstrates that a single dose of HU induces testicular germ cell apoptosis dose dependently in a time- and stage-specific pattern and that spermatogonia and early spermatocytes are the main target cells of induced apoptosis. It appears that apoptosis of male germ cells is a common response to testicular toxicants, and the death of these germ cell serves to eliminate the affected cells preventing fertility disturbance and protecting future generations from the transmission of damaged DNA. ACKNOWLEDGMENTS We gratefully acknowledge Dr. E. M. Eddy, NIEHS/NIH, USA, for kindly providing HSP70-2 antibody. We thank Dr. M. Smith for his helpful comments and critical reading of this manuscript. This study was financially supported by grants from the Japanese Ministry of Education, Science, Sports and Culture. J.S. is a recipient of scholarships awarded by the Korea Science and Engineering Foundation (KOSEF) and the Kyoto University Foundation.

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