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Unité de Gamétogenèse et Génotoxicité, Institut National de la Santé et de la Recherche Médicale (INSERM) Unité 566, Commissariat à l’Energie Atomique, Université Paris 7–Denis Diderot (G.D., C.L., C.P., C.R., C.D., R.H.), 92265 Fontenay-aux-Roses; and Institut de Génétique et de Biologie Moléculaire et Cellulaire, INSERM Unité 184, Centre National de la Recherche Scientifique/INSERM/Université Louis Pasteur, Collège de France (A.K.), BP 163, 37404 Illkirch-Cedex, France
Address all correspondence and requests for reprints to: Dr. Christine Levacher, Institut National de la Santé et de la Recherche Médicale Unité 566, Commissariat à l’Energie Atomique, Université Paris 7–Denis Diderot, DSV/DRR/SEGG/LDRG, Bâtiment 5A, RDC, Route du Panorama, 92265 Fontenay-aux-Roses, France. E-mail: christine.levacher{at}cea.fr.
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gene was inactivated. The homozygous inactivation of ERß (ERß–/–) increased the number of gonocytes by 50% in 2- and 6-d-old neonates. The numbers of Sertoli and Leydig cells and the level of testicular testosterone production were unaffected, suggesting that estrogens act directly on the gonocytes. The increase in the number of gonocytes did not occur during fetal life but instead occurred just after birth, when gonocytes resumed mitosis and apoptosis. It seems to result from a decrease in the apoptosis rate evaluated by the terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling method and cleaved caspase-3 immunohistochemical detection. Last, mice heterozygous for the ERß gene inactivation behaved similarly to their ERß–/– littermates in terms of the number of gonocytes, apoptosis, and mitosis, suggesting that these cells are highly sensitive to the binding of estrogens to ERß. ER inactivation had no effect on the number of neonatal gonocytes and Sertoli cells. In conclusion, this study provides the first demonstration that endogenous estrogens can physiologically inhibit germ cell growth in the male. This finding may have important implications concerning the potential action of environmental estrogens. |
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and ERß, are widely distributed in the testes of various species, including mice (2, 3). Studies reporting poor semen quality in some male patients with a mutation in the ER gene (4) or with an aromatase deficiency (5, 6) or reporting infertility in transgenic mice with deletions of the gene encoding ER (ERKO) (7) or aromatase (Cyp19; ArKO) (8, 9) have also indicated that estrogens are essential for testicular function. In humans, a decrease in sperm count (10, 11) and an increase in the incidences of testicular cancer (12), cryptorchidism, and hypospadia (13) have been observed in many countries in the last 50 yr. Such male reproductive disorders in humans and in wildlife have been attributed to the increase in concentration of xenobiotics, and of xenoestrogens in particular, in the environment and in food (14, 15). This hypothesis was first put forward in 1993 by Sharpe and Skakkebaek (16), who developed an "estrogen hypothesis" in which they argued that estrogen-like molecules could alter adult male fertility by acting early in gonad development and that inappropriate exposure to estrogen during fetal or neonatal life could affect adult reproductive function. Indeed, the male offspring of women treated with diethylstilbestrol (DES), a potent synthetic estrogen, during pregnancy have been reported to show a high incidence of reproductive disorders, such as atrophic testes and sperm abnormalities (17, 18). Furthermore, the exposure of laboratory animals during fetal or neonatal life to high doses of estradiol or DES leads to various abnormalities of the testis and reproductive tract (13), including low testis weight and germ cell volume/testis (19, 20), an increase in germ cell apoptosis (21), an increase in the incidence of tumors of the reproductive tract (22), and a decrease in sperm fertilizing ability (23). Adverse short-term effects of early transplacental exposure to high doses of DES, such as accelerated testicular development, abnormal differentiation of gonocytes and Sertoli cells, fetal Leydig cell hyperplasia in mice (24), and changes in steroidogenic factor-1 gene expression in rats (25) have also been observed before birth. Estrogens have also been shown to inhibit the development of gonocytes and Leydig and Sertoli cells in rat fetal testis in vitro (26) and to act directly on the testis to exert their deleterious effects. Last, a positive direct effect of estrogens on gonocyte development has been evidenced in purified gonocyte cultures (27).
Thus, estrogens have been shown to affect all cell types within the testis and may alter the genetic cascade ensuring normal testis development. However, although most of these deleterious effects are obvious in experiments with pharmacological doses of estrogens, no evidence has yet been presented that low doses equivalent to the level of human exposure to environmental estrogens have any effect. A few studies have reported the effects of exposure to low doses of estrogens. They have shown that neonatal exposure to low doses of DES has no long-term adverse effect in terms of testis size or fertility but may induce short-term stimulatory effects on the first wave of spermatogenesis (28). Fielden et al. (29) observed that the fetal and lactational exposure of mice to genistein at levels similar to those present in the human diet significantly increased in vitro fertilizing ability of epididymal sperm.
Nonetheless, the physiological role of estrogens in the development of male germ cells is unclear because the cellular and molecular mechanisms by which estrogens could control testis development and functions are unknown. We tested the hypothesis that endogenous estrogens affect early testis development by investigating gametogenesis during fetal and neonatal life in mice lacking ERs (ERKO and ERßKO mice) (30). Overall testicular development appears to be normal in ERKO mice (31), but fetal and neonatal testicular development has never been analyzed in these animals. Studies with these mice should also make it possible to discriminate between the roles of ER and ERß in the control of early testicular development and gametogenesis by endogenous estrogens.
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Materials and Methods |
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(ER–/–) or ERß (ERß–/–) were produced by Dupont et al. (30) (exon 3 of these genes, encoding the first zinc finger of the DNA-binding domain was targeted for the disruption) and generously provided by Pierre Chambon (Institut de Génétique et de Biologie Moléculaire et Cellulaire, Illkirch, France). Mice were housed under controlled photoperiod conditions (lights on 0700–2000 h) and were supplied with commercial feed and tap water ad libitum. Heterozygous males were caged with heterozygous females for the night, and the day after an overnight mating was counted as d 0.5 post conception (0.5 dpc). Natural birth occurred on fetal d 19.5, which was counted as d 0 postpartum (0 dpp). Pregnant mice were anesthetized on gestational d 13.5, 15.5, or 17.5 by ip injection of 4 mg/100 g sodium pentobarbital (Sanofi, Libourne, France), and the fetuses were quickly removed from the uterus. Fetuses were dissected under a binocular microscope, their sex was determined based on the morphology of the gonads, and the testes were collected from male fetuses. Male neonates were killed by decapitation on postnatal d 2 and 6, and their testes were removed immediately. All the animals were genotyped by PCR of biopsy DNA, as described previously (30). All animal studies were conducted in accordance with the NIH Guide for Care and Use of Laboratory Animals.
Testicular treatment
The testes were fixed in Bouin’s fluid for 2 h and embedded in paraffin. The high density of the cells in the 13.5 dpc testis made it necessary to cut 3-µm sections, whereas for all other stages we cut 5-µm sections.
Measurement of testicular volume
One in 20 sections of testes removed from 2 or 6 dpp neonates was mounted on slides, deparaffinized, rehydrated, and stained with Tushmann’s blue. We measured the areas of each section with a computerized video densitometer (Histolab; Microvision Instruments, Evry, France) and added them together. The resulting value was multiplied by 20 and by the section thickness (5 µm) to obtain the testicular volume.
Testicular cell counting
Gonocytes.
Serial sections were mounted on slides, deparaffinized, rehydrated, and stained with Tushmann’s blue. On fetal d 13.5 and 6 dpp, identification of gonocytes was not easy with this staining. Therefore, we performed immunostaining for AMH, the gonocytes being identified as the cells that remained unstained (32) (Fig. 1A
, B, K, L). AMH immunostaining was performed with the Vectastain Elite ABC kit (Vector Laboratories, Burlingame, CA). Briefly, sections were incubated successively with 3% H2O2 in distilled water for 10 min to inactivate endogenous peroxidases and with a mouse anti-AMH polyclonal antibody (1:250), generously provided by N. Di Clemente (INSERM Unité 493, Clamart, France), for 1 h. This primary antibody was detected with a biotinylated goat antirabbit secondary antibody and the avidin-biotin-peroxidase complex. Peroxidase activity was visualized using 3,3'-diaminobenzidine as the substrate. We used morphological criteria for the identification of gonocytes on fetal d 15.5 (Fig. 1, C and D) and 17.5 (Fig. 1, E and F) and on neonatal d 2 (Fig. 1, G and H) (33, 34). The accuracy of the counting method was assessed on the basis of two criteria. First, the minimum number of sections to be counted was selected such that the calculated number of total cells was similar to that obtained by counting a greater number of sections. Second, we checked that similar numbers of gonocytes were found in each testis from the same mouse fetus. We achieved this by mounting one in 10 sections for fetal d 13.5 and one in 20 sections for other stages, which means that the gonocytes were counted in a minimum of nine sections equidistantly distributed along the testis. The sum of the values for one testis was multiplied by 10 or 20 to obtain the crude count (CC) per testis. The Abercrombie formula (35) was used to correct for double counting resulting from the appearance of a single cell in two successive sections: TC = CC x S/(S + D), where TC is the true count, S is the section thickness (3 or 5 µm), and D is the true mean diameter of the cell nuclei. D equals the mean of the nuclear diameters measured (DM) on the section divided by 
/4 to correct for the overexpression of smaller profiles in sections through spherical particles. DM was measured on each testis by means of at least 100 random determinations with a computerized video micrometer (Microvision Instruments). At 6 dpp, because of the great number of germ cells, we counted all these cells only in the middle section. This number was divided by the area of the section to obtain the density of germ cells per square millimeter. All counts were done blind.
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, G and H) and counted (36). All counts were done blind.
Leydig cells.
The Leydig cells in the testes of mice killed on postnatal d 2 were identified by immunocytochemical detection of 3ß-hydroxysteroid dehydrogenase, using an antibody provided by Dr. G. Defaye (INSERM Unité 244, Grenoble, France) (Fig. 1, I and J). Immunostaining was performed with the Vectastain Elite ABC kit (Vector Laboratories), as described previously (36). Leydig cells were counted on one section in 20. All counts were done blind.
Measurement of bromodeoxyuridine incorporation index
The neonates (2 dpp) were given ip injections of 1 ml/100 g 5-bromo-2'-deoxyuridine (BrdU) and 5-fluoro-2'deoxyuridine solution, according to the manufacturer’s recommendations (Cell Proliferation kit; Amersham, Buckinghamshire, UK), 3 h before they were killed. BrdU incorporation into proliferating cells was detected by immunocytochemistry, as described previously (37). The BrdU incorporation index (percentage of labeled gonocytes) was obtained by counting at least 500 gonocytes on the sections.
Terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick end labeling (TUNEL) staining
Apoptotic cells were detected in situ using the TUNEL method (37), with the modifications described previously (38). We mounted each fifth section on a single slide, washed that slide twice in PBS, and heated it in permeabilization solution (0.05 M Tris; pH 10.6) in a microwave oven for 3 min at 750 W plus 7 min at 370 W. Endogenous peroxidase activity and nonspecific protein binding were blocked by incubation for 10 min in 3% hydrogen peroxide. The slides were then washed three times, for 5 min each, in PBS. Sections were incubated for 30 min with 3% BSA and 20% normal bovine serum (Sigma, St. Louis, MO) in PBS for saturation purposes and were then covered with the TUNEL mix (Roche, Mannheim, Germany), which contained calf thymus TdT, fluorescein-dUTP, and unlabeled dNTP in reaction buffer [200 mM potassium cacodylate, 25 mM Tris-HCl, 1 mM cobalt chloride, and 0.25 mg/ml BSA (pH 6.6)], and incubated for 1 h in a humidified chamber at 37 C. The reaction was stopped by washing the slides twice, for 5 min each, in PBS and then twice, for 5 min each, in Tris (pH 7.4). Sections were then blocked again by incubating for 30 min at room temperature in 3% BSA, 20% normal sheep serum (Sigma), and 1% blocking reagent (Roche) in Tris (pH 7.4). The sections were treated with a peroxidase-labeled fluorescein sheep Fab antibody (Roche) for 30 min in a humidified chamber at 37 C. The slides were washed in PBS, and the peroxidase was visualized using 3,3'-diaminobenzidine as the substrate. Positive controls were treated with DNaseI (100 µg/ml; Sigma) for 10 min at room temperature beforehand to induce DNA fragmentation in all the nuclei, and negative controls were incubated without TdT. TUNEL-positive gonocytes were counted on every section, and the number obtained was multiplied by five to obtain the total number of TUNEL-positive gonocytes per testis.
Immunohistochemical staining for cleaved caspase-3
Because cleaved caspase-3 is involved in most of the apoptotic pathways, we chose its detection as a marker of apoptosis (39). One in five sections of testis removed from 2 dpp neonates was mounted on a single slide, washed twice in PBS, and heated in permeabilization solution (0.05 M Tris; pH 10.6) in a microwave oven for 3 min at 750 W plus 7 min at 370 W. Endogenous peroxidase activity and nonspecific protein binding were blocked by incubation for 10 min in 3% hydrogen peroxide. The slides were then washed three times, for 5 min each, in PBS and incubated for 1 h with normal goat serum and with the cleaved caspase-3 antibody (1:100e, 9661; Cell Signaling Technology, Beverly, MA) (39) for the next 2 h. The distribution of the primary antibody was revealed with a biotinylated goat antirabbit secondary antibody and an avidin-biotin-peroxydase complex (Vectastain Elite ABC kit; Vector Laboratories). Peroxidase was visualized with 3–3'-diaminobenzydine tetrahydrochloride. Stained gonocytes were counted on every section, and the number obtained was multiplied by five to obtain the total number of cleaved caspase-3-positive gonocytes per testis.
Testosterone secretion
Immediately after their removal, testes taken from 2-d-old mice were incubated on Millipore (Bedford, MA) filters (pore size, 0.45 µm) for 3 h, as described previously (40). Briefly, each testis was cut into three pieces and placed on a filter floating on 0.4 ml Ham’s F-12/DMEM (1:1; Life Technologies, Inc., Grand Island, NY) in tissue culture dishes incubated at 37 C, in an humidified atmosphere containing 95% air and 5% CO2. For each animal, one testis was incubated with basal medium and the other with medium containing 100 ng/ml ovine LH (National Institutes of Health, Bethesda, MD; LH S19; 1.01 U/mg). The testosterone secreted into the medium was measured by RIA, as described previously (41).
Estradiol assay
For the determination of testicular estradiol content, 20 testes from 2 dpp wild-type mice were homogenized. Steroids were extracted with diethylether as described previously. Estradiol was measured by RIA using a specific antibody from Biosys (Compiegne, France) (42). The sensitivity was 4–5 pg/tube. The intraassay and interassay coefficients of variation were 5 and 9%, respectively.
Statistical analysis
The results are presented as means ± SEM. The statistical significance of the difference between the mean values for two different genotypes was evaluated using Student’s unpaired t test. One-way ANOVA was used for the simultaneous comparison of data from more than two groups. The 
2 test was used to compare observed and expected percentages.
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or ERß gene–/– or ERß–/–) were phenotypically normal at birth. On postnatal d 2, the sex ratio in litters was not significantly different from the expected 50%, and the mutant allele was transmitted to the offspring with the expected Mendelian inheritance pattern (data not shown). At 2 and 6 dpp, the studied mutations affected neither the weight of the pups nor their testis volume (Tables 1 and 2).
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or ERß gene–/– and ERß–/– mice appeared normal in terms of overall histology and similar to that of wild-type littermates. The mean diameter of gonocyte nuclei was not affected by inactivation of the ER genes [8.13 ± 0.07 µm in wild-type mice (n = 13), 8.16 ± 0.16 µm in ERß–/– mice (n = 13), and 8.01 ± 0.21 µm in ER–/– mice (n = 4)]. Two days after birth, ER–/– neonates and wild-type neonates had similar numbers of gonocytes (Fig. 2). In contrast, inactivation of the ERß gene resulted in a significant increase in the number of gonocytes (52% more in ERß+/– and 49% more in ERß–/– mice than in wild-type mice; Fig. 2). At 6 dpp, the density of gonocytes per square millimeter was significantly increased in ERß–/– mice compared with the wild-type mice. Because the testis volume was unchanged, this suggests that the total number of gonocytes per testis is still increased 6 d after birth in ERß–/– mice (Table 2).
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). The diameter of gonocyte nuclei was not affected by ERß gene inactivation at any of the ages studied (data not shown). In the wild-type population, the number of gonocytes per testis quadrupled from 13.5 to 15.5 dpc and remained stable thereafter. The number of gonocytes was unaffected by inactivation of the ERß gene until 17.5 dpc. Thereafter, in contrast to what was observed in the wild type, the number of gonocytes in ERß–/– mice increased and was significantly higher in these mice than in wild-type mice at 2 d of age [11,187 ± 539 gonocytes in 17.5 dpc fetuses (n = 4) and 17,459 ± 811 in 2 dpp neonates (n = 13); P < 0.001]. The increase in the number of gonocytes was similar in ERß+/– and ERß–/– mice [11,424 ± 1115 gonocytes in 17.5 dpc fetuses (n = 4) and 17,829 ± 1,917 in 2 dpp neonates (n = 7); P < 0.05].
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), at which time the gonocytes had resumed mitosis, as 13.28 ± 2.4% (n = 9) were in S phase. The BrdU incorporation index (Fig. 4B) in ERß+/– and ERß–/– mice was not found to differ significantly from that in wild-type mice, due to the considerable variability in this variable. However, if we pooled animals homozygous and heterozygous for the mutation, the difference between this group and the controls was statistically significant.
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, C–F). By immunofluorescent staining, we observed a colocalization of the TUNEL signal with the cleaved caspase-3 (data not shown). Inactivation of the ERß gene resulted in a significant decrease in the number of TUNEL-positive gonocytes per testis in both ERß+/– and ERß–/– neonates [858 ± 81 in wild-type (n = 5) vs. 514 ± 127 in ERß+/– (n = 4), P < 0.05, and 662 ± 19 in ERß–/– (n = 5), P < 0.05]. The same result was found for the number of cleaved caspase-3-positive gonocytes per testis [611 ± 69 in wild-type (n = 4) vs. 311 ± 19 in ERß+/– (n = 4), P < 0.01, and 355 ± 56 in ERß–/– (n = 4), P < 0.05]. When expressed as a percentage of the total number of gonocytes per testis, there were at least 50% fewer TUNEL- or cleaved caspase-3-positive gonocytes in ERß+/– and ERß–/– mice than in the wild type (Fig. 4, D and F).
Number of Sertoli cells in neonates with inactivation of the ER or ERß gene
The number of Sertoli cells at 2 dpp was unchanged after ER or ERß inactivation compared with their wild-type littermates (Fig. 5).
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).
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).
Testicular estrogen content
In wild-type 2 dpp mice, the testicular content in estradiol was 0.48 pg/testis, which corresponds to 4 nM.
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Discussion |
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Our study also demonstrated that estrogens exert their effects on male fetal/neonatal gametogenesis via ERß. Indeed, the homologous recombination used to disrupt ERß in the ERßKO mice (30) resulted in a complete elimination of ERß protein, and so the possibility of an alternative ERß active variant is excluded. In contrast, ER does not seem to be involved because its inactivation had no effect on germ cell development. However, the implication of the membrane ERs have already been described in other tissues (43) and cannot be excluded here because the ERKO model does not allow us to consider it. The effects of estrogens on germ cell development described here are unlikely to result from an estrogenic action on the hypothalamus-pituitary gonadotropic system, although little is known about gonadotropin levels in ERßKO. Basal testicular testosterone production was unaffected in ERß–/– neonates, which suggests that the secretion of LH and GnRH is normal. Furthermore, no change in the number of gonocytes has been observed in vivo in fetal rats after decapitation or in mice with inactivation of the FSH receptor (44) (Racine, C., S. Migrenne, E. Moreau, and R. Habert, unpublished data) or in vitro after treatment of rat fetal testis by gonadotropins (45). Thus, estrogens seem to act directly on the testis, raising important questions as to whether they act directly on the germ cells or indirectly, via another cell type. Because the number and activity of Leydig cells were unaffected by ERß inactivation, these cells are unlikely to be involved in changes in the number of germ cells. Furthermore, it has been suggested that estrogeninduced changes in fetal Sertoli cell proliferation are an essential determinant in the estrogen-induced decrease in germ cell content (19, 21). In studies with an organ culture system, we recently found that Sertoli cells did not mediate the effect of estrogens on gonocyte development: 1) estrogens decreased the number of gonocytes 2 d before the number of Sertoli cells decreased; 2) the function of Sertoli cells appeared to be unaffected; and 3) an antiestrogen blocked the effect of estrogens on gonocyte development without affecting Sertoli cell proliferation (26). The data presented here support the hypothesis that estrogens act directly on germ cells because ERß inactivation had no effect on the number of Sertoli cells. Moreover, our observation of a biological effect of ERß inactivation and no effect of ER inactivation is consistent with observations that ERß is present (46) in the gonocytes of the fetal/neonatal mouse, whereas ER is not (47).
In this study, we also established the ontogenesis of the ERß-mediated inhibitory effect of endogenous estrogens on gonocyte development in the mouse. We began by establishing the pattern of change in the number of gonocytes during normal mouse fetal development, which had never been reported before. We found that the number of gonocytes increased until 15.5 dpc and remained constant thereafter. This is consistent with the well known quiescent phase of gonocytes, in which no mitotic or apoptotic activity occurs and which extends from 15.5 dpc until birth in mice (34, 48, 49). This also indicates that after birth, mitosis and apoptosis resume (34, 49) and are perfectly balanced such that the number of cells remains stable, at least until 2 dpp. We found that 13% of gonocytes were BrdU positive, whereas only 2% were apoptotic, due to the much longer duration of the S phase (few hours) than of DNA fragmentation (some minutes) (45, 50, 51). In ERß–/– mice, no change in gonocyte development was observed until 17.5 dpc. This suggests that the migration and proliferation of primordial germ cells, which occur before 11.5 dpc (52), and the proliferation of fetal gonocytes until 15.5 dpc are estrogen-independent processes. The lack of effect of ERß deficiency during this early developmental period appears to conflict with our previous data showing that high doses of 17ß-estradiol or DES decreased the number of gonocytes in 14.5 dpc rat testes in culture (26). This difference in results may be due to differences in experimental approaches (in vivo vs. in vitro, deficiency in a physiological context vs. addition of high concentrations) or to species differences because, in the mouse, ERß mRNA is detectable from 14.5 dpc but the protein does not become detectable until 16.5 dpc (i.e. after the end of the proliferative period in fetal gonocytes) (46), whereas in the rat, ERß protein is present on fetal d 16.5, at a time when proliferation is continuing in this species (53). Our study suggests that estrogens do not affect the number of gonocytes during the quiescent period, in line with previous reports in the rat for other controlling factors (TGFß1, TGFß2, retinoic acid) (36, 37). However, 2 dpp ERß–/– neonates had 50% more germ cells than their wild-type littermates. This is essentially due to a lower rate of apoptosis and, to a lesser degree, to a higher rate of mitosis. This is consistent with previously described changes in the apoptotic and mitotic activities of gonocytes in response to in vitro estrogen treatment (26). Nevertheless, the effects of ERß deficiency observed during neonatal life may also originate from an impairment of gonocyte maturation occurring during the fetal quiescent period. Indeed, it has been clearly shown that a deleterious stress (irradiation) applied during the quiescent period does not change the number of gonocytes during this period but increases the rate of apoptosis and inhibits the mitosis of gonocytes during neonatal life (54).
These findings are important not just for endocrinology but also with respect to their implications concerning the possible involvement of environmental xenoestrogens in the reported decrease in male fertility (3, 13). Previous studies have shown that exposure to estrogens in perinatal life inhibits spermatogenesis, but only for high doses of these molecules (13). We show here that the concentrations of estrogens present in the fetal and neonatal testis are sufficient to inhibit germ cell development during perinatal life. The reason for this discrepancy is not clear. It probably results from differences in the experimental approaches used because our study involved the creation of conditions equivalent to estrogen deficiency, whereas the other studies dealt with exogenous exposure to estrogenic compounds. Because the aromatase gene is expressed in fetal and neonatal Sertoli cells (55), the local testicular production of estrogens may minimize the effects of exogenous estrogens. Indeed, testicular 17ß-estradiol content is approximately 4 nM (0.48 pg/mouse testis) at 2 dpp. Moreover, the results for heterozygous neonates are striking. Heterozygous animals are generally considered to have the wild-type phenotype or to have less marked abnormalities than homozygous animals. For instance, males and females heterozygous for the ER deletion are fertile whereas homozygotes are sterile (56). In this study, heterozygous neonates behaved like ERßKO littermates, despite having half the normal amount of ERß, suggesting that germ cells may be highly sensitive to estrogens during fetal/neonatal life. Thus, the contrast between extreme experimental conditions, strong excess, and deficiency in estrogens indicates that current knowledge of the action of estrogens is insufficient to reconcile these two sets of findings, particularly because some studies have also showed a positive effect of estradiol on spermatogenesis in various species (27, 57, 58, 59, 60).
In conclusion, our study demonstrates an ERß-mediated in vivo inhibitory effect of endogenous estrogens on establishment of the male germ cell lineage throughout fetal/neonatal life in the mouse. This effect results from an increase of gonocyte apoptosis. These findings provide important insight into the poorly explored field of fetal and neonatal gametogenesis. They also provide support for the hypothesis that exposure to environmental xenoestrogens during fetal/neonatal testicular development may have deleterious effects on male fertility in adulthood.
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Acknowledgments |
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Footnotes |
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Abbreviations: AMH, Anti-Müllerian hormone; BrdU, 5-bromo-2'-deoxyuridine; DES, diethylstilbestrol; dpc, day(s) post conception; dpp, day(s) postpartum; ER, estrogen receptor; TdT, terminal deoxynucleotidyl transferase; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling.
Received October 31, 2003.
Accepted for publication March 15, 2004.
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References |
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(ER) and ß (ERß) on mouse reproductive phenotypes. Development 127:4277–4291[Abstract]
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and ß in neonatal rat testis: identification of gonocytes as targets of estrogen exposure. Biol Reprod 68:867–880This article has been cited by other articles:
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C. R. Cederroth, O. Schaad, P. Descombes, P. Chambon, J.-D. Vassalli, and S. Nef Estrogen Receptor {alpha} Is a Major Contributor to Estrogen-Mediated Fetal Testis Dysgenesis and Cryptorchidism Endocrinology, November 1, 2007; 148(11): 5507 - 5519. [Abstract] [Full Text] [PDF]
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G. Delbes, C. Duquenne, J. Szenker, J. Taccoen, R. Habert, and C. Levacher Developmental Changes in Testicular Sensitivity to Estrogens throughout Fetal and Neonatal Life Toxicol. Sci., September 1, 2007; 99(1): 234 - 243. [Abstract] [Full Text] [PDF]
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M L Gould, P R Hurst, and H D Nicholson The effects of oestrogen receptors {alpha} and {beta} on testicular cell number and steroidogenesis in mice Reproduction, August 1, 2007; 134(2): 271 - 279. [Abstract] [Full Text] [PDF]
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 K. Boukari, M. L. Ciampi, A. Guiochon-Mantel, J. Young, M. Lombes, and G. Meduri Human fetal testis: source of estrogen and target of estrogen action Hum. Reprod., July 1, 2007; 22(7): 1885 - 1892. [Abstract] [Full Text] [PDF]
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 J. Merlet, C. Racine, E. Moreau, S. G. Moreno, and R. Habert Male fetal germ cells are targets for androgens that physiologically inhibit their proliferation PNAS, February 27, 2007; 104(9): 3615 - 3620. [Abstract] [Full Text] [PDF]
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 C. Kress, C. Gautier-Courteille, H. B. Osborne, C. Babinet, and L. Paillard Inactivation of CUG-BP1/CELF1 Causes Growth, Viability, and Spermatogenesis Defects in Mice Mol. Cell. Biol., February 1, 2007; 27(3): 1146 - 1157. [Abstract] [Full Text] [PDF]
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S. Assinder, R. Davis, M. Fenwick, and A. Glover Adult-only exposure of male rats to a diet of high phytoestrogen content increases apoptosis of meiotic and post-meiotic germ cells Reproduction, January 1, 2007; 133(1): 11 - 19. [Abstract] [Full Text] [PDF]
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G. Delbes, C. Levacher, and R. Habert Estrogen effects on fetal and neonatal testicular development. Reproduction, October 1, 2006; 132(4): 527 - 538. [Abstract] [Full Text] [PDF]
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 C Roger, S Lambard, A Bouskine, B Mograbi, D Chevallier, M Nebout, G Pointis, S Carreau, and P Fenichel Estrogen-induced growth inhibition of human seminoma cells expressing estrogen receptor {beta} and aromatase J. Mol. Endocrinol., August 1, 2005; 35(1): 191 - 199. [Abstract] [Full Text] [PDF]
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J. A. Arreguin-Arevalo and T. M. Nett A Nongenomic Action of 17{beta}-Estradiol as the Mechanism Underlying the Acute Suppression of Secretion of Luteinizing Hormone Biol Reprod, July 1, 2005; 73(1): 115 - 122. [Abstract] [Full Text] [PDF]
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G. Delbes, C. Levacher, C. Duquenne, C. Racine, P. Pakarinen, and R. Habert Endogenous Estrogens Inhibit Mouse Fetal Leydig Cell Development via Estrogen Receptor {alpha} Endocrinology, May 1, 2005; 146(5): 2454 - 2461. [Abstract] [Full Text] [PDF]
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