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Endocrinology Vol. 144, No. 6 2617-2622
Copyright © 2003 by The Endocrine Society

Pituitary Hormones Inhibit the Function and Differentiation of Fetal Sertoli Cells

Stéphanie Migrenne, Chrystèle Racine, Florian Guillou and René Habert

Functional Differentiation of Gonads Laboratory, Gametogenesis and Genotoxicity Unit (S.M., C.R., R.H.), Institut National de la Santé et de la Recherche Médicale U566-Commissariat á l’Énergie Atomique-Université Paris 7, 92265 Fontenay aux Roses, France; and Station de Physiologie de la Reproduction des Mammifères Domestiques (F.G.), Institut National de la Recherche Agronomique/Centre National de la Recherche Scientifique, Unité de Recherche Associée 1291, 37380 Nouzilly, France

Address all correspondence and requests for reprints to: Prof. R. Habert, Unité Gamétogenèse et Génotoxicité, Institut National de la Santé et de la Recherche Médicale U566-Commissariat á l’Énergie Atomique-Université Paris 7, Département de Radiobiologie et Radioprotection, Bat 5, BP6, Route du Panorama, 92265 Fontenay aux Roses Cédex, France. E-mail: rene.habert{at}cea.fr.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Although the role of pituitary hormones in fetal Sertoli cell proliferation is well understood, their involvement in fetal Sertoli cell differentiation is poorly documented. In this study, we evaluated rat fetal Sertoli cell function by measuring basal transferrin secretion ex vivo and transferrin and anti-Müllerian hormone (AMH) mRNA levels in vivo. The differentiation state of the Sertoli cells was estimated from the amount of transferrin secreted ex vivo after acute stimulation with FSH. Surprisingly, we found that the amount of transferrin secreted by each Sertoli cell in basal condition and after acute FSH stimulation decreased between 18.5 and 21.5 day post coitum (dpc), which corresponds to the onset of pituitary hormone secretion. All of the Sertoli cell parameters measured (basal and FSH-stimulated transferrin secretion ex vivo, transferrin and AMH mRNA levels in vivo) were higher in 21.5-dpc fetuses that had been decapitated on 16.5 dpc than in control littermates. Furthermore, immunostaining for AMH was strongly increased after decapitation. Taken together, these results suggest that pituitary hormones in the fetus and in the immature or adult rat differently regulate Sertoli cells, which suggests that fetal Sertoli cells have their own particular physiology.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
SERTOLI CELLS IN the testes of adult rats are of major functional importance because they provide the structural support necessary for germ cell maturation and create the optimal environment for spermatogenesis. Sertoli cells start to carry out these functions in the fetus, because gonocytes isolated in vitro die rapidly (1, 2, 3). Furthermore, the number of spermatozoa formed by the adult testis depends on the number of Sertoli cells (4), which is in turn determined during fetal and postnatal life.

In the rat, the Sertoli cell precursors present in seminiferous cords by 13.5 days post coitum (dpc) are dividing. The percentage of proliferating Sertoli cells peaks at 20.5 dpc and gradually declines during the first 2 or 3 wk of postnatal life (5, 6). Sertoli cell proliferation is regulated by a number of pituitary factors that are first expressed between 16.5 and 19.5 dpc, depending on the factors considered (7, 8, 9, 10, 11). FSH plays a central role in regulating Sertoli cell proliferation in the fetus. Specific suppression of fetal FSH dramatically reduces the percentage of rat Sertoli cells entering the S phase of the cell cycle (12). Similarly, the number of Sertoli cells in the testes of newborn hypogonadal (hpg) mice, which have low levels of circulating gonadotropins, is 30% lower than in normal newborn mice (13). In the same way, FSH receptor-deficient or FSHß-deficient mice models show a reduction of testis weight and tubule diameter, suggesting a decrease in Sertoli cell number (14, 15, 16, 17). The data concerning the role of TSH are more controversial, because some authors reported a decrease in Sertoli cell proliferation in neonatal rats with induced hypothyroidism and others reported an increase (18, 19). Testosterone may also be involved in Sertoli cell proliferation because androgen treatment reverses the increase in Sertoli cell mitosis induced by hemicastration of immature rats (20).

Contrary to the regulation of Sertoli cell proliferation, the role of pituitary hormones in the regulation of fetal and neonatal Sertoli cell function and differentiation is poorly documented. All available data concern the ability of these cells to respond to a pituitary hormone. In neonatal Sertoli cell cultures, the addition of FSH and/or T3 alters the expression of anti-Müllerian hormone (AMH), inhibin {alpha}, and aromatase (21, 22, 23). The injection of FSH into rat fetuses reduces both AMH mRNA and protein levels (24). However, no data are currently available concerning the physiological role of the total pituitary secretions in the regulation of fetal Sertoli cell function and differentiation.

The objective of this study was to determine the overall role of the pituitary hormones in fetal Sertoli cell differentiation. Accordingly, we decapitated fetal rats in utero at 16.5 dpc. Thus, the whole pituitary gland was removed before the onset of hormonal secretion. The specific function of Sertoli cells was evaluated by studying the expression of the fetal Sertoli cell differentiation markers, AMH and transferrin.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Wistar strain rats from Iffa Credo (L’Arbresle, France) were housed under a controlled photoperiod with lighting from 0700–1900 h, fed a commercial diet (UAR, Villemoisson sur Orge, France), and allowed tap water ad libitum. Males were caged with the females at night. Because the estimated time of ovulation was 0200 h, the day after an overnight mating was counted as d 0.5 post conception. Natural birth occurred between d 21.5 at 1400 h and d 22.5 at 1800 h. All animal studies were conducted in accordance with the Guides for Care and Use of Laboratory Animals (NIH Guide).

Incubations of testes ex vivo
Pregnant females were anesthetized between 0900 and 1300 h on 16.5, 18.5, and 21.5 dpc by an ip injection of sodium pentobarbital (Sanofi Pharmaceuticals, Inc. Libourne, France; 4 mg/100 g body weight), and the testes of male fetuses were removed under a stereological microscope. To obtain pieces of similar size, testes were cut into 2, 4, or 16 pieces at 16.5, 18.5, and 21.5 dpc, respectively. These pieces were rinsed three times for 20 min to eliminate any contamination by transferrin secreted by the liver, placed in tissue culture dishes, and incubated for 3 h at 37 C in 0.4 ml of culture medium in a humidified atmosphere containing 95% air/5% CO2, as previously described (25). The culture medium was Ham’s F12/DMEM (1:1) (Life Technologies, Inc., Grand Island, NY) containing 15 mM HEPES, 7.5% sodium bicarbonate, 0.35% glutamine (Flow Laboratories, Inc., McLean, VA), and 80 µg/ml gentamicin (Gentalline, Schering-Plough Corp., Levallois-Perret, France). The effect of FSH was evaluated by comparing one testis cultured in medium containing 200 mU/ml recombinant FSH (rFSH; a generous gift of 12,000 IU/mg from Dr. B. Mannaerts, Organon International, Oss, The Netherlands), with the other testis from the same fetus cultured in medium alone (control). Note that this dose corresponds to approximately 10 times the concentration of serum FSH in newborn control mice (Migrenne, S., E. Moreau, A. Dierich, P. Pakarinen, R. Habert, and C. Racine, unpublished data).

After incubation, testes were stored at -20 C in RNA Plus reagent (Bioprobe Systems, Montreuil-sous-Bois, France) for RNA extraction and subsequent RT-PCR analysis. The incubation media were used for transferrin RIA.

Surgical procedures
Between 0900 and 1300 h on 16.5 dpc, pregnant rats were anesthetized by ip injection of 10.5 mg ketamine per 100 g body weight (Imalgène 500; Rhône Mérieux, Lyon, France). One fetus in each uterine horn was then decapitated in utero by the method of Jost (26). The mothers were again anesthetized by sodium pentobarbital between 0900 and 1300 h on 21.5 dpc for collection of fetuses. Only 26% of the fetuses decapitated on 16.5 dpc survived. The undecapitated fetuses served as sham controls.

Immediately after removal, fetal testes were processed in one of four ways: incubated ex vivo for measurement of the basal and FSH-stimulated transferrin production, fixed in Bouin’s fluid for evaluation of Sertoli cell development, stored at -20 C for assay of protein content, or stored in RNA Plus reagent for RNA extraction and subsequent RT-PCR analysis.

Morphometric analysis of fetal Sertoli cells
The number of fetal Sertoli cells in testes from control fetuses removed at 16.5, 18.5, or 21.5 dpc and from fetuses decapitated at 16.5 dpc and removed at 21.5 dpc were counted on histological sections according to previously described morphological criteria (27).

Immunohistochemical staining for AMH
Immunostaining for AMH was performed as previously described, using anti-AMH antibodies, kindly given by B. Vigier (Unité de Biologie du Développement et Biotechnologies, Jouy en Josas, France) (27). Six tissue blocks from decapitated fetuses and their control littermates were used, and three to six sections from each block were immunostained.

RNA extraction and analysis by RT-PCR
RT-PCR was used to study gene expression in rat fetal testes, because of the small amount of RNA available. Total RNA was extracted from whole testes using the RNA Plus kit (Bioprobe Systems). RNAs were quantified by measuring absorbance at 260 nm, and their quality was checked by gel electrophoresis. PCR primer pairs (Table 1Go and Refs. 28, 29, 30) were selected from different exons of the corresponding genes to distinguish any PCR products arising from possible chromosomal DNA contaminants. One microgram of total testicular RNA was reverse transcribed, and target transferrin or AMH cDNA and ß-actin cDNA (used as internal control) were coamplified by PCR as previously described (31), except that transferrin was annealed at 60 C and AMH at 56 C. For transferrin ontogenesis, PCR analysis was performed without an internal control, under the usual PCR conditions (32). RT-PCR products were analyzed in 2% agarose gel (wt/vol) stained with ethidium bromide. DNA bands were photographed and scanned before semiquantitative analysis with NIH Image 1.62 software for Macintosh. Lastly, the densitometric signal was expressed as a percentage of the value obtained for a normal littermate.


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  		Table 1. Primer pairs used for RT-PCR analysis of gene expression in fetal rat testis

 
Assays
Transferrin was assayed in the culture media of fetal testes by RIA according to Le Magueresse et al. (33). For this purpose, standard rat transferrin was purchased from Sigma (St. Louis, MO), and the usable range of the assay was 0.2–200 ng/tube, with an intrasample coefficient of variation of 8%.

Statistical analysis
All values are means ± SEM. The significance of the differences between the mean values for treated and control testes in vitro was evaluated by Student’s paired t test. Data for decapitated and control fetuses were compared by Student’s unpaired t test. Testicular protein contents were compared by the Mann-Whitney U test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effects of decapitation on body and testis growth and on the number of Sertoli cells in male fetuses are shown in Table 2Go. The headless body weight of rat fetuses increased 4.3-fold between 16.5 and 18.5 dpc and 3.8-fold between 18.5 and 21.5 dpc. After decapitation, body weight was 13.7% lower than that of the control littermates.


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  		Table 2. Effects of decapitation on body and testis growth and on the number of Sertoli cells in male fetuses

 
In control fetuses, the testicular protein content increased 2.4-fold between 16.5 and 18.5 dpc and 2.7-fold between 18.5 and 21.5 dpc. Decapitation led to a 10.5% decrease compared with control littermates.

In control fetuses, the number of Sertoli cells increased 2.7-fold between 16.5 and 18.5 dpc and 11.6-fold between 18.5 and 21.5 dpc. At 21.5 dpc, the testes of fetuses decapitated 5 d earlier were macroscopically indistinguishable from those of control littermates. The number of fetal Sertoli cells diminished by 22% after decapitation, but the mean diameter of their nuclei was not significantly different in decapitated and control littermates (6.70 ± 0.18 µm, n = 6; vs. 6.80 ± 0.18 µm, n = 6, respectively).

Expression of transferrin in the control fetal testis
To determine whether transferrin is expressed in rat fetal testis, we used RT-PCR to analyze transferrin mRNA levels from 14.5–21.5 dpc. Transferrin mRNA was detected in fetal testis at the earliest age studied (14.5 dpc, 25 PCR cycles) and in many other fetal tissues such as the ovary (Fig. 1Go). The amount of basal transferrin secreted by the whole testis did not change between 16.5 and 18.5 dpc but significantly increased between 18.5 and 21.5 dpc (Fig. 2AGo). The addition of FSH to testes ex vivo increased both the amount of transferrin mRNA and the amount of transferrin secreted (Fig. 2Go, A and B). Because Sertoli cells are the only cell type in the testis known to possess FSH receptors (34), this increase implies that transferrin is secreted by fetal Sertoli cells, in agreement with the data reported for the pubertal and adult testis (35).



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  		Figure 1. RT PCR analysis of transferrin mRNA in fetal rat testes (A) and various fetal and postnatal rat tissues (B). Total RNA was extracted from fetus testes at 14.5–21.5 dpc (A, lanes 1–8) and from various rat tissue samples [B, lane 1, liver (16.5 dpc); lane 2, testis (14.5 dpc); lane 3, ovary (14.5 dpc); lane 4, spleen (14 dpp); lane 5, muscle (16.5 dpc); lane 6, brain (16.5 dpc); and lane 7, tongue (14 dpp)]. RNAs were then analyzed by RT-PCR using transferrin-specific primers. Results are given for three representative independent determinations. MW, Molecular weight.

 


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  		Figure 2. Effect of FSH on the amount of transferrin secreted ex vivo by the fetal rat testis (A), the transferrin mRNA level in vivo (B), and the amount of transferrin secreted per fetal Sertoli cell (C). Fetal testes were removed at 16.5, 18.5, and 21.5 dpc, washed three times for 20 min to eliminate serum transferrin, and incubated for 3 h with (FSH) or without (basal) 200 mU/ml rFSH. A. The amount of transferrin secreted into the medium was measured by RIA. Values are means ± SEM of 6–10 determinations. B, After incubation, total RNA was extracted from control testes (basal) and FSH-stimulated testes (FSH) and subjected to RT-PCR analysis. Transferrin mRNA levels in basal and FSH-stimulated testes were normalized compared with ß-actin. Gels are representative of three independent determinations. C, The amount of transferrin secreted into the medium (presented in A) was divided by the number of Sertoli cells per testis. Values are means ± SEM of three to six determinations. *, P < 0.05; **, P < 0.01; and ***, P < 0.001 according to the Student’s t test.

 
Interestingly, the amount of transferrin secreted per testicular Sertoli cell did not change between 16.5 and 18.5 dpc, but dropped dramatically between 18.5 and 21.5 dpc, with the capacity to respond to FSH stimulation decreased (+691% at 18.5 dpc and +169% at 21.5 dpc; P < 0.05; Fig. 2CGo).

Effect of decapitation on the expression of transferrin by fetal Sertoli cells
We measured the amount of transferrin secreted by the fetal testis after 3 h in the presence or absence of FSH. The amount of transferrin secreted was expressed per testis and per Sertoli cell (Fig. 3Go). Decapitation did not change the amount of transferrin secreted by 21.5 dpc testes ex vivo in either condition when values were expressed per testis (Fig. 3AGo). However, both basal and FSH-stimulated transferrin secretions were greater in the testes of decapitated fetuses than in those of their control littermates when values were expressed per Sertoli cell (Fig. 3BGo).



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  		Figure 3. Effects of fetal decapitation on basal and FSH-stimulated transferrin secretion. Fetuses were decapitated in utero at 16.5 dpc. At 21.5 dpc, testes were removed from decapitated living fetuses and from their sham control littermates and washed three times for 20 min to eliminate serum transferrin. Testes were then incubated for 3 h with (FSH) or without (basal) 200 mU/ml rFSH. The amount of transferrin secreted into the medium was measured by RIA (A) and divided by the number of Sertoli cells in each testis (B). Results are means ± SEM of six determinations. *, P < 0.05; **, P < 0.01; and ***, P < 0.001 according to the Student’s t test.

 
Furthermore, the testicular transferrin mRNA level was 1.7-fold higher in 21.5 dpc decapitated fetuses than in control littermates (Fig. 4Go).



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  		Figure 4. RT-PCR analysis of transferrin and AMH mRNA after fetal decapitation. Fetuses were decapitated in utero at 16.5 dpc. At 21.5 dpc, testes were removed from decapitated living fetuses (D) and from their sham control littermates (C). Total RNA was extracted and analyzed by RT-PCR. Transferrin and AMH levels in the testes of decapitated and control fetuses were normalized according to ß-actin. Results are expressed as means ± SEM (n = 5) of the percentage of expression relative to a normal littermate (control) analyzed on the same gel. *, P < 0.05; and **, P < 0.01 using the Student’s t test.

 
Effect of decapitation on AMH expression
Semiquantitative RT-PCR analysis showed that the AMH mRNA level was 2.2-fold higher in decapitated fetuses than in control littermates (Fig. 4Go). Immunostaining of AMH protein was also markedly higher in decapitated fetuses (Fig. 5Go).



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  		Figure 5. Effect of decapitation on the expression of AMH. A, Testis from control fetus; B, testis from decapitated fetus. Fetuses were decapitated in utero at 16.5 dpc. At 21.5 dpc, testes were removed from decapitated living fetuses (B) and from their sham control littermates (A), fixed in Bouin’s fluid, immunostained for AMH, and counterstained with hematoxylin. Note the increased staining after decapitation. Scale bars, 10 µm.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
These results demonstrate that during late fetal life, pituitary hormones regulate Sertoli cell proliferation positively and affect their function and differentiation negatively.

This study was motivated by our previous results obtained using rat fetuses that had been decapitated in utero, which showed that fetal Leydig cell activity depends on LH, whereas Leydig cell differentiation is LH-independent (36).

The same approach was used to investigate the physiological relevance of the cumulative effects of pituitary hormones in vivo. The rat is a highly suitable model for this purpose because the rat placenta does not produce chorionic gonadotropin (37, 38, 39) and because the gonadotropins produced by the pregnant female do not penetrate the fetus (40). Rats were decapitated at 16.5 dpc, i.e. before the onset of any pituitary hormone expression (7, 8, 9, 10, 11), which made it possible to remove all gonadotropins and other pituitary hormones from the fetal plasma.

Decapitation significantly reduced the number of Sertoli cells during late fetal life. This finding is consistent with the report by Orth (12) showing that late decapitation of rat fetuses or injection of FSH antiserum reduces the amount of 3H-thymidine incorporated into Sertoli cells. Furthermore, the number of Sertoli cells decreases in hpg mouse testes during the perinatal period (13, 41) but significantly increases when intact neonatal rats and neonatal hpg mice are treated with FSH (41). It is established that testosterone stimulates Sertoli cell proliferation in immature rat testes (13, 41), and we previously showed that decapitated fetuses have a very low testicular testosterone content (36). However, the reduction in the number of Sertoli cells is probably not directly linked to this decrease in testosterone, because no androgen receptors are detected in Sertoli cells before 5 days postpartum (dpp; Ref. 42). Nevertheless, androgen receptors have been immunolocated in fetal Leydig and peritubular cells (43), and therefore we cannot rule out the possibility that androgen acts through these cell types.

The important finding of this investigation was that the expression of AMH is physiologically regulated by the cumulative action of hypophyseal hormones. AMH is a perfect marker for determining the state of fetal Sertoli cell differentiation because its expression drops sharply during the postnatal period. After fetal decapitation, AMH mRNA and protein levels both rose significantly. These results are in accordance with those of in vitro studies showing that AMH expression decreases after the addition of T3 and/or FSH to postnatal rat testes in culture (23), or after injecting FSH into rat fetuses (24). In the same way, the injection of newborn animals with FSH decreases AMH immunohistochemical staining on the second postnatal day. However, this treatment has no further effect on the fourth postnatal day (44). Furthermore, the injection of FSH from birth to the sixth postnatal day increases AMH levels determined at the seventh postnatal day (45). Taken together, these results suggest that FSH inhibits AMH expression during late fetal and early postnatal life and stimulates its expression thereafter.

It is well established that testosterone decreases AMH synthesis at puberty (46). However, during fetal life, the increase in AMH expression observed here after decapitation was probably not the result of the decrease in testosterone production (8, 36), because the injection of neonates with LH does not modify AMH protein or mRNA levels (44) and there is no evidence that androgen receptors are present in fetal Sertoli cells (42), as mentioned above.

In immature and adult rat testes, transferrin is a major product secreted by differentiated Sertoli cells (35) and is acutely stimulated by FSH. It is therefore a useful marker of Sertoli cell function and differentiation (47, 48). Transferrin mRNA has been detected in the rat testis as early as 5 dpp (49). Here, we provided the first evidence that transferrin mRNA is present in the rat fetal testis as early as 14.5 dpc. Gelly et al. (50) immunodetected transferrin in the rat fetal testis at 13.5 dpc, but exclusively in the gonocytes. This location is probably consecutive to the endocytosis of transferrin arising from the Sertoli cells or from the plasma by germ cells. In humans, transferrin mRNA has been detected in 8- to 12-wk-old fetal testes after 20 d of culture (51). We demonstrated for the first time that transferrin secretion is acutely stimulated by FSH from 16.5 dpc onward, the earliest time point we examined. This is consistent with previous data showing that FSH receptors are expressed and functional in the rat testis as early as 15.5 dpc (52, 53). The fact that the amount of transferrin secreted by the fetal testis increased after acute FSH stimulation allowed us to use it as a fetal Sertoli cell marker. We provide two pieces of evidence that the expression of transferrin is subject to negative physiological control by pituitary hormones during fetal life. First, when basal testicular transferrin secretion ex vivo (an index of its in vivo secretion) was divided by the number of Sertoli cells per testis, we found a surprising decrease between 18.5 and 21.5 dpc, i.e. at the onset of pituitary hormone secretion. Second, after fetal decapitation, both the transferrin mRNA level and the basal secretion per Sertoli cell rose. These findings are interesting because these two parameters drop dramatically in hypophysectomized adult rats (54). Furthermore, acute FSH-stimulated transferrin secretion per fetal Sertoli cell (an index of cell differentiation in vivo) also dropped when hypophyseal secretion started in controls, and fetal decapitation increased this secretion. Conversely, Sertoli cells of the adult rat do not increase the acute FSH-stimulated transferrin secretion after hypophysectomy (54). Taken together, these results suggest, for the first time, that Sertoli cell functions (as reflected by basal secretion ex vivo and the mRNA level in vivo) and differentiation (as indicated by acute FSH-stimulated secretion ex vivo) are differently regulated by pituitary hormones in the fetus and in immature or adult rats. This suggests the existence of a shift from negative to positive control between the fetal and immature states. Further experiments will be necessary to determine when this shift actually occurs and to establish whether it is linked to the fact that fetal Sertoli cells proliferate, whereas immature and adult cells do not.

In conclusion, our data suggest that pituitary hormones directly or indirectly regulate Sertoli cell function and differentiation in the fetus. Further studies using transgenic mice are currently being performed in our laboratory to identify the pituitary hormone responsible for this regulation. Contrary to the situation in the adult testis, pituitary hormones in the fetal testis physiologically inhibit both function and differentiation state in each Sertoli cell. This suggests that, like Leydig cells (36, 55), Sertoli cells have their own particular physiology during fetal life.


   Acknowledgments
 
We thank B. Vigier (Unité de Biologie du Développement et Biotechnologies, Jouy en Josas, France) for the generous gift of anti-AMH antibodies. We are also grateful to Monique Faro for technical assistance.


   Footnotes
 
This work was supported by the Institut National de la Santé et de la Recherche Médicale, Commissariat á l’Énergie Atomique, Université Paris 7, and the Fondation pour la Recherche Médicale. S.M. was the recipient of fellowships from the Ministère de la Recherche et de la Technologie and from Fondation pour la Recherche Médicale.

Abbreviations: AMH, Anti-Müllerian hormone; dpc, day post coitum; dpp, days postpartum; hpg, hypogonadal; rFSH, recombinant FSH.

Received November 5, 2002.

Accepted for publication February 5, 2003.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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