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Endocrinology, doi:10.1210/en.2003-0072
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Endocrinology Vol. 144, No. 8 3651-3662
Copyright © 2003 by The Endocrine Society

Unaltered Development of the Initial Follicular Waves and Normal Pubertal Onset in Female Rats after Neonatal Deletion of the Follicular Reserve

Céline J. Guigon, Séverine Mazaud, Maguelone G. Forest, Sylvie Brailly-Tabard, Noëlline Coudouel and Solange Magre

Laboratoire de Physiologie et Physiopathologie, Centre National de la Recherche Scientifique, Unité Mixte de Recherche 7079, Université Pierre et Marie Curie (C.J.G., S.M., N.C., S.M.), 75005 Paris, France; Institut National de la Santé et de la Recherche Médicale (INSERM), Unité (U)-329, Laboratoire de Pathologie Hormonale Moléculaire, Hôpital Debrousse (M.G.F.), 69322 Lyon, France; and INSERM, U-135, Laboratoire d’Hormonologie et Biologie Moléculaire, Hôpital de Bicêtre (S.B.-T.), 94275 Le Kremlin Bicêtre, France

Address all correspondence and requests for reprints to: Dr. S. Magre, Laboratoire de Physiologie et Physiologie, Unité Mixte de Recherche 7079, Université Paris VI, 7 quai St. Bernard, 75005 Paris, France. E-mail: solange.magre{at}snv.jussieu.fr.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In rats, the pool of primordial follicles is established within the first 3 d postnatally (dpn). Immediately after their differentiation, a subset of follicles begins to grow and constitutes the initial follicular waves. In this study we investigated the development of these early growing follicles after deletion of the primordial follicle pool induced by 1.5 Gy {gamma}-irradiation at 5 dpn. Within only 24 h, i.e. at 6 dpn, 99% of the primordial follicles disappeared, whereas most of the growing follicles remained unaffected. The study of these surviving follicles throughout the immature period has shown that their subsequent growth proceeded normally, as assessed by proliferating cell nuclear antigen immunostaining and follicular counts. No modification in the process of follicular atresia, studied by terminal deoxynucleotidyltransferase-mediated deoxy-UTP-fluorescein nick end labeling and Southern blot of DNA fragmentation analysis, was observed. Complementary analysis, by either in situ hybridization for inhibin subunits, P450 aromatase, and LH receptor mRNAs or plasma dosages of 17ß-estradiol and inhibin B, further showed that follicular maturation was unaltered. In line with these observations, pubertal onset was normal, regarding both age and ovulation rate. Nevertheless, as a consequence of the nonrenewal of the growing pool, the follicular complement was practically exhausted at puberty, and 90% of the females evidenced sterility by 4 months. Altogether, our results demonstrate that the deletion of the primordial follicle pool has induced no modification in the growth pattern of the early growing follicles that develop as their counterparts in control ovaries. Within the immature period, the initial follicular waves ensure the ovarian functionality and thus play a key role in the initiation of reproductive life.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
IN MAMMALS, FOLLICULOGENESIS begins with the fragmentation of ovigerous cords in small follicular units, the primordial follicles, each unit consisting of a single layer of flattened pregranulosa cells surrounding a quiescent oocyte (1, 2). As the pool of primordial follicles is nonrenewable, ovarian life span is tightly limited by the size of the follicle stockpile; the dwindling supply or eventual exhaustion of the primordial follicle pool is referred to as reproductive senescence in mammals, more commonly termed menopause in women. In rats, as soon as the primordial follicles are formed, a subset of follicles located in the core of the ovary begins to grow. These initial follicle waves develop through primary, preantral follicle stages and reach the antral stage by 3 wk of life (2). During this period, both the number of primordial follicles recruited into the growing follicle pool and the rate of follicular growth are higher than those in adult cycling animals (3). Most of the antral follicles developing in the immature period are destined to be eliminated from the ovary because, unlike those of adult females, no cyclic gonadotropin surge can rescue them from the process of follicular atresia (4). Among the initial waves of follicular growth, a few follicles nonetheless reach the preovulatory stage and ovulate around the fifth week of life, heralding the establishment of the estrous cycling. Thus, already in immature life a subtle balance between recruitment of primordial follicles and follicular growth, on the one hand, and follicular atresia, on the other, is required for the production of ova at the initiation of reproductive life.

Numerous experimental animal models have been used to investigate the consequences of modifications of the size of the primordial follicle pool. Mice deficient for the proapoptotic gene bcl-2-associated X protein (Bax) maintain a larger resting pool of primordial follicles in adulthood and possess numerous growing follicles in the ovary at an advanced chronological age (5). Conversely, zinc finger X (Zfx) mutant mice and XO mice, which are endowed with few primordial follicles at birth, show diminished fertility and a shortened reproductive life span (6, 7). The size of the follicle stockpile, however, is unlikely to be the only parameter determining ovarian life span. It has been reported in several models of dysgenesic ovaries that the reduction of the follicular stockpile could result in an alteration of follicular dynamics. In rats treated in utero with busulfan, a chemical that destroys germ cells, the primordial follicle endowment at birth is greatly reduced, and the rate of early follicular growth is increased in the immature period. Young adult females exhibit a normal number of large antral follicles until exhaustion of the follicular stockpile (8). Recent results from our group have shown that in rats irradiated in utero and lacking nearly 99% of primordial follicles, the growth of the first follicular waves is delayed, and follicular atresia is reduced. Normal numbers of ova are shed at the onset of puberty, and females are fully fertile for a couple of months (9). Thus, modifications in the pattern of follicular growth and possibly in the process of follicular atresia could occur to regulate the whole follicular complement when the primordial follicle pool is absent, as early as during the immature period.

In experimental models of dysgenesic ovaries, the reduced size of the follicle stockpile results either from a failure of germ cell development or a reduction in the number of germ cells during fetal life (6, 10, 11, 12), that is, before the initiation of folliculogenesis. To our knowledge, there are no data concerning the consequences of the depletion of the primordial follicle pool soon after its differentiation, i.e. when the first follicular waves have already initiated their growth. The aim of the present work has been to study the effect of the elimination in the neonatal period of the primordial follicle pool on subsequent ovarian development in the rat. In rodents, oocytes from the resting pool are known to be very sensitive to irradiation (13, 14). To drastically deplete the population of primordial follicles, whole body {gamma}-irradiation was performed at 5 d postnatally (dpn) when the radiosensitivity of the rat ovary was maximal (15).

After ascertaining the effect of the irradiation on follicular complement, we carried out, in parallel to the study on fertility, analysis of ovarian differentiation by in situ approaches and dosages of plasma ovarian hormones. From 6 dpn until puberty, we conducted quantitative and qualitative studies of follicular development using follicular counts, in situ hybridization of follicular maturation markers, and analysis of follicular atresia.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals and irradiation
Pregnant Sprague Dawley female rats were purchased from Charles River Laboratories, Inc. (L’Arbresle, France). After birth (0 dpn), female pups were grouped by 8–10/mother. At 5 dpn, they were whole body exposed to {gamma}-irradiation using a 60Co source with a total dose of 1.5 Gy at a dose rate of 0.25 Gy/min. From 21 dpn, four or five weaned females were housed in a single cage and weighed daily. From 30 dpn, females were checked daily for the appearance of vaginal opening, the external criteria of pubertal onset (16). Animals were maintained in standard lighting (12 h of light, 12 h of darkness) and temperature (22 C), with food and water ad libitum.

Tissue collection
Ovaries were collected daily from 5–12 dpn and then at 15, 21, 28, and 32 dpn, on the day of vaginal opening and between 2–6 months. For animals studied at 32 dpn, only nonpubertal females were used. On the day of vaginal opening, histological examination of the ovary was used to check the presence of corpora lutea (first estrus). In both groups, 70% of the females were at first estrus, and the others were at first proestrus. The ovaries were either snap-frozen in liquid nitrogen and stored at -80 C or processed for further morphological studies. From 21 dpn, ovaries, uteri, and pituitary were weighed.

At 21, 28, and 32 dpn and on the day of puberty, females were killed by decapitation, and trunk blood was collected with Pasteur pipettes containing heparin. Animals were always killed in the afternoon (1500–1700 h). Blood samples were centrifuged at 3000 x g for 15 min, and the plasma collected was stored at -20 C until assayed.

Tissue processing
For histological examination of ovarian morphology and follicle counts, ovaries were placed in Bouin’s fixative for at least 24 h, dehydrated in alcohol, and paraffin-embedded using standard protocols. Sections of 5 µm thickness were mounted on glass slides and stained with hematoxylin and eosin. For in situ hybridization, terminal deoxynucleotidyltransferase-mediated deoxy-UTP-fluorescein nick end labeling (TUNEL) assay, and immunohistochemistry, ovaries were fixed in 2% paraformaldehyde-PBS (pH 7.2) for 1–3 h depending on organ size. After washing in PBS and in increased concentrations of sucrose in PBS, ovaries were embedded in Tissue-Tek OCT compound (Miles, Inc., Elkhart, IN) and kept at -80 C. Frozen sections of 5 µm thickness were mounted onto 3-aminopropyltriethoxysilane (Sigma-Aldrich Corp., St. Louis, MO)-treated glass slides and stored at -20 C.

Quantitative assessment of oocytes, follicles, and corpora lutea
Quantification of oocytes at 6 dpn and follicles in older animals (15 dpn to first estrus) was carried out as previously described (9) with some modifications (Table 1Go) using the oocyte nucleus as a marker. At 6 dpn up to the large primary stage, oocytes were counted in every second section, whereas oocytes in preantral follicles were counted in every fifth section to avoid several counts of the same oocyte. In older animals, follicle or corpora lutea counts were performed in every fifth section. Oocytes were deemed unhealthy if they were convoluted and condensed or fragmented. A follicle was considered atretic if it contained a degenerating oocyte and/or at least two pycnotic granulosa cells.


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  		TABLE 1. Classification of oocytes and follicles, modified from Ref. 9

 
In situ hybridization
The cDNAs used for the riboprobe synthesis were obtained after RT-PCR performed on rat ovaries with appropriate oligonucleotide primers (Table 2Go) and ligation into PGEM-T Easy vectors (Promega Corp., Madison, WI). The cDNA clones were verified after sequencing. Sense and antisense riboprobes were generated by transcription with digoxigenin-labeled deoxy-UTP (Roche, Mannheim, Germany) and the appropriate T7 or SP6 polymerase (Roche).


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  		TABLE 2. In situ hybridization probes

 
After thawing, sections were briefly treated with chloroform to remove lipids and rehydrated in PBS. Tissue sections were fixed in 2% paraformaldehyde-PBS (pH 9) for 20 min, rinsed in PBS, and incubated for 10 min in 0.1 M triethanolamine (pH 8) with 0.25% acetic anhydride (Sigma-Aldrich Corp.). Prehybridation was carried out for 2 h at 55 C in 50% deionized formamide, 2x sodium saline citrate (SSC), 100 µg/ml yeast tRNA, 4 mM EDTA (pH 8), 5x Denhardt’s, 2.5% dextran sulfate, and 250 µg/ml salmon testis DNA (Sigma-Aldrich Corp.). Hybridization was carried out overnight at 55 C with riboprobes diluted in prehybridation mix without EDTA and salmon testis DNA. Treatment of the sections with ribonuclease A in 500 mM NaCl, 10 mM Tris (pH 7.5), and 5 mM EDTA for 30 min at 37 C was followed by a high-stringency wash in 30% formamide/0.1x SSC at 65 C for 1 h. After a brief wash in T1 buffer [100 mM Tris (pH 7.5) and 150 mM NaCl] and a 1-h wash in 5% blocking reagent (Roche) in T1, sections were briefly rinsed. They were then incubated for 1.5 h with the antidigoxigenin alkaline phosphatase-conjugated antibody (dilution, 1:500; Roche). Sections were incubated overnight with the chromogen substrates of alkaline phosphatase, 5-bromo-4-chloro-3-indolyl-phosphate (0.175 mg/ml) and nitro blue tetrazolium (0.337 mg/ml), in the presence of levamisole (24 µg/ml) in 100 mM Tris (pH 9.5), 100 mM NaCl, and 50 mM MgCl2 (Roche).

Southern blot analysis of DNA fragmentation
In three independent experiments, genomic DNA was extracted as previously described (17) from ovaries at 21, 28, and 32 dpn and on the day of first estrus. In each experiment a single ovary (32 dpn and first estrus) and a pool of two ovaries (21 and 28 dpn) were homogenized in a homogenization buffer [100 mM NaCl, 10 mM EDTA, 300 mM Tris-HCl, and 200 mM sucrose (pH 8)] with 0.6% sodium dodecyl sulfate and 60 µg proteinase K. Homogenates were incubated for 4 h at 62–64 C and gently mixed by repeated passages through a pipette. After the addition of 0.7 M potassium acetate (pH 5.3) to facilitate protein precipitation, lysates were centrifuged at 5000 x g for 15 min at 4 C. After collection of the supernatant and digestion of RNA with ribonuclease deoxyribonuclease-free (Roche), DNA was phenol/chloroform-extracted and quantified by spectrophotometry. A total of 5 µg DNA was 3'-end-labeled with 20 µM digoxygenin-11-dideoxy-UTP (Roche) and 20 U terminal deoxynucleotidyltransferase (Roche) in terminal deoxynucleotidyltransferase reaction buffer with 200 mM cacodylate for 1 h at 37 C. Digoxigenin-labeled samples were resolved through 2% agarose gels, transferred to a positively charged nylon membrane (Roche) as previously described (18), and covalently attached using UV cross-linking. After washing and blocking for 30 min, the membrane was incubated for 30 min with an antidigoxigenin alkaline-phosphatase-conjugated antibody (dilution, 1/20,000; Roche). The 3'-end-labeled DNA was detected by chemiluminescence after incubation of the membrane with CDP-Star (Roche) for 5 min and exposition to Hyperfilm ECL (Amersham International, Little Chalfont, UK). Low molecular weight (<1.2 kb) DNA fragments were quantified using NIH Image (1.62) analysis software. Each independent experiment was performed in triplicate. Quantification of DNA fragmentation was presented as relative to the amount of DNA fragmentation in control ovaries at 6 dpn.

TUNEL
After thawing and rehydration, sections were postfixed in 4% paraformaldehyde-PBS (pH 7.2), rinsed in PBS, and incubated for 1 h at room temperature with a mix containing fluorescein-deoxy-UTP (TUNEL Label, Roche) and terminal deoxynucleotidyltransferase (TUNEL enzyme, Roche). Detection of apoptotic cells was occasionally performed on sections previously processed for in situ hybridization. In that case, sections were directly rinsed in PBS and incubated similarly for 1 h with the reaction mix. Cells exhibiting nuclear staining were considered apoptotic. Negative controls, lacking the labeling enzyme, yielded no reaction product (data not shown).

Proliferating cell nuclear antigen (PCNA) immunohistochemistry
After thawing, sections were delipidized in chloroform and rehydrated in PBS. The endogenous activity of the peroxidase enzyme was blocked by treatment in 3% H2O2 for 10 min. Sections were incubated for 1 h at room temperature with the primary antibody (dilution, 1:400 in 0.5% BSA; sc-56, Santa Cruz Biotechnology, Inc., Santa Cruz, CA) and thereafter for 1 h with an antimouse biotinylated antibody (dilution, 1:200 in 0.5% BSA; RPN 1001, Amersham Biosciences) and for 30 min with a peroxidase-conjugated streptavidin-horseradish complex (LSAB+ Kit, DAKO Corp., Carpinteria, CA). The reaction product was developed using 3,3'-diaminobenzidine tetrahydrochloride (LSAB+ Kit, DAKO Corp.). For negative controls (not shown), primary antibody was omitted.

Hormone assays
Individual plasma were assayed, except at 21 and 28 dpn for steroid measurements, for which plasma from two or three animals were pooled. Plasma 17ß-estradiol (E2) and testosterone (T) were assayed by specific RIAs after ethyl ether extraction and chromatographic purification on Celite columns as previously described (19, 20). The plasma concentration of inhibin B was determined using a two-site ELISA specific for each peptide (Oxford Bioinnovation, Oxon, UK) as previously described (21). Intraassay coefficients of variation were 4.9% and 5%, and interassay coefficients of variation were 12% and 9% for inhibin B and steroids, respectively. The sensitivities of the assays were 10 pg/ml, 3 pg/tube, and 5 pg/tube for inhibin B, E2, and T, respectively.

Fertility assessment
Irradiated and control females (n = 11 in each group) were continuously mated from 60 dpn on. One male was housed with a group of three or four females. Pregnant females at 14–15 d gestation were isolated. Newborn pups were counted and kept with their mother until 2 dpn to check breastfeeding and eventual lethality. The mother was further isolated for 1 wk before new mating. At the end of the mating experiment, embryonic implantation sites in the uterus were counted under a dissection microscope.

Statistical analysis
Data collected from at least three different animals were analyzed using GBStat 6.0 software and were expressed as the mean ± SEM. Statistical significance was determined by one- or two-way ANOVA in conjunction with a post hoc multiple comparison test (Scheffé’s test) if necessary. Transformation of data in log values was sometimes required when high differences in means between the two groups resulted in nonproportional variances. To compare the percentage of pregnant females between the two groups, {chi}2 tests were performed.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
The irradiated females presented neither signs of developmental abnormalities nor disease consecutive to the whole body exposure to 1.5 Gy at 5 dpn. No significant differences were noted in body weights between irradiated and control females for both immature and adult animals (Table 3Go and data not shown). Weights of ovaries, uterus, and pituitary were similar between control and irradiated females throughout the immature period and on the day of puberty (Table 3Go). The onset of puberty, assessed by the appearance of vaginal opening, occurred at the same age in the two groups (33.6 ± 0.3 dpn in controls vs. 34.4 ± 0.4 dpn in irradiated females; n = 25 and n = 31, respectively; P = 0.46).


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  		TABLE 3. Body, ovarian, uterus, and pituitary weights in control (ctr) and irradiated (irr) females

 
Deletion in oocytes of the follicle stockpile
At the time of treatment, i.e. 5 dpn, the ovary contained resting oocytes (naked oocytes and oocytes in primordial follicles; Table 1Go) located in the cortex as well as growing oocytes in primary and preantral follicles, distributed mainly in the core of the ovary (data not shown). Similar organization was observed at 6 dpn in control ovaries (Fig. 1Go, Aa and Ac). In irradiated females, growing oocytes were clearly visible in the core of the ovaries (Fig. 1AGob). Very few healthy resting oocytes were present, however, in the cortex (Fig. 1AGod). Follicular counts showed that the pool of healthy resting oocytes was suppressed by more than 99% in irradiated ovaries (17.2 ± 2.1 vs. 2821.2 ± 170.6 in controls; P < 0.001; Fig. 1BGoa). Small primary follicles were also affected by the treatment, as shown by the attrition of numerous germ cells (Fig. 1Go, Ad and Bb) and the reduction by 35% in the number of healthy oocytes within this follicle category (302.6 ± 20.1 vs. 458.8 ± 15.75 in controls; P < 0.05; Fig. 1BGoa). In contrast, oocytes in large primary and preantral follicles were unaffected by the treatment (Fig. 1Go, Aa–Ad). No difference was observed in the number of healthy and unhealthy oocytes in either category between irradiated and control ovaries (Fig. 1Go, Ba and Bb).



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  		FIG. 1. Effect of irradiation on the ovary 24 h after exposure. A, Histological sections of control (Aa and Ac) and irradiated (Ab and Ad) ovaries. Control and irradiated ovaries exhibit similar organization, with growing oocytes (GO) located in the center of the ovary (Aa and Ab). Compared with controls, irradiated ovaries have few, if any, resting oocytes (RO) in the cortex (Ac and Ad). Many oocytes in small primary follicles are degenerating in irradiated ovaries (white arrowheads; Ad). Note the presence of primordial follicle-like structures that do not contain an oocyte in the irradiated cortex (black arrows). Scale bar, 100 µm. B, Numbers of healthy (Ba) and unhealthy (Bb) oocytes in control ({blacksquare}) and irradiated ({square}) ovaries. Oocytes were counted on histological sections as described in Materials and Methods. Data are the mean ± SEM of five ovaries per group. *, P < 0.01; **, P < 0.001. C, In situ end labeling of apoptotic cells on frozen sections of control (Ca) and irradiated (Cb and Cc) ovaries. In control ovaries, very few apototic cells are detected at 6 dpn (Ca). In irradiated ovaries, apoptotic cells are numerous at 6 dpn (24 h after irradiation; Cb), but become rare at 7 dpn (48 h post irradiation; Cc). Scale bar, 100 µm.

 
Interestingly, when compared with control ovaries, irradiated ovaries displayed a similar number of degenerating oocytes at the resting stage (632.6 ± 90.9 vs. 684.2 ± 82.1 in controls; P > 0.05) and a higher number of degenerating oocytes in small primary follicles (169.4 ± 8.4 vs. 8.6 ± 2.9; P < 0.001; Fig. 1BGob). Thus, 24 h after irradiation, most of the resting oocytes had already been destroyed by irradiation, whereas oocytes in small primary follicles were still dying. To clarify whether the treatment could destroy oocytes contained in follicles beyond the small primary follicle stage later than 24 h, we compared germ cell apoptosis at 6 and 7 dpn by TUNEL assay. At 6 dpn, irradiated females displayed numerous TUNEL-positive cells in the ovarian cortex compared with controls, whereas at 7 dpn rare TUNEL-positive cells were observed in the irradiated ovaries (Fig. 1Go, Ca–Cc). Thus, the irradiation massively affected the follicular content of the ovary within 24 h through its deleterious effect on germ cells of the primordial follicle pool. In contrast, the treatment only slightly affected follicles belonging to the initial follicular waves.

Follicular growth until puberty
To determine whether the proliferative capacity of granulosa cells and thus the subsequent development of the growing follicles were modified in irradiated ovaries, an immunohistochemical analysis of PCNA (22) was carried out at 6 dpn. A comparable PCNA immunostaining was observed in the nucleus of granulosa cells in primary and preantral follicles in control and irradiated ovaries (Fig. 2Go, A and B). In the following days, the development of growing follicles proceeded similarly in irradiated and control ovaries. As early as 8 dpn, both control and irradiated females displayed follicles at the antral stage (data not shown). At 15 and 21 dpn, the numbers of preantral and antral follicles were similar in control and irradiated ovaries (Fig. 3Go, A and B).



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  		FIG. 2. Immunolocalization of PCNA on frozen sections of control and irradiated ovaries 24 h after irradiation. In both control (A) and irradiated (B) ovaries, numerous granulosa cells have a strong PCNA nuclear staining in growing follicles (black arrowhead). Note that growing oocytes (white arrow) and a few cells in the mesenchymal compartment (black arrow) also stain positively. Scale bar, 100 µm.

 


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  		FIG. 3. Evolution of the population of healthy preantral (A) and antral (B) follicles from 15 dpn to first estrus in control ({blacksquare}) and irradiated ({square}) ovaries. Follicle counts were performed on histological sections as described in Materials and Methods. Bars represent the mean ± SEM of three ovaries at 15 dpn and four ovaries at the other ages. Data were analyzed by two-way ANOVA. A: Effect of the treatment, P < 0.0001; effect of age, P < 0.0001; treatment by age, P = 0.01. B: Effect of the treatment, P < 0.001; effect of age, P < 0.0001; treatment by age, P = 0.0006. *, P < 0.05; **, P < 0.01.

 
During the immature period, however, irradiated ovaries displayed a sequential reduction in the number of the different classes of growing follicles. At 15 dpn, the population of primary follicles was reduced by 95% in irradiated ovaries (17.3 ± 4.7 vs. 355 ± 34.15; P < 0.001; n = 3 in each group). At 28 dpn, the number of preantral follicles (Fig. 3AGo) was reduced by 73% (49.75 ± 3.0 vs. 184.5 ± 19.1 in controls; P < 0.001) and reached very low values on the day of first estrus (8 ± 3 vs. 138.2 ± 15.2 in controls; P < 0.001). The population of healthy antral follicles (Fig. 3BGo) was decreased by 59% at 32 dpn (29.5 ± 2.9 vs. 71.25 ± 7.9 in controls; P < 0.05) and by about 84% on the day of first estrus (12.5 ± 3.7 vs. 77.2 ± 3.5 in controls; P < 0.001). Nevertheless, at first estrus the ovaries exhibited a similar number of fresh corpora lutea in both groups (4.2 ± 0.48 in irradiated females vs. 5.00 ± 0.58 in controls; n = 5 and n = 4, respectively; P = 0.33).

Follicular atresia
To study the process of follicular atresia during the immature period, we performed TUNEL assay (Fig. 4AGo), analysis of DNA fragmentation after gel electrophoresis (Fig. 4BGo), and follicle counts (Fig. 4CGo). The first TUNEL-positive granulosa cells were detected at 12 dpn in preantral follicles and at 15 dpn in antral follicles in both control and irradiated ovaries (data not shown). From 21 dpn, most of the antral follicles displayed numerous TUNEL-positive granulosa cells in the two groups (Fig. 4Go, Aa and Ab). Analysis of DNA fragmentation from 21 dpn to first estrus after gel electrophoresis revealed a laddering pattern, with DNA fragments differing in size by 180–200 bp (Fig. 4BGoa). In both groups quantification of DNA fragmentation showed that the apoptotic process was higher in the immature period than at first estrus. No significant difference between control and irradiated ovaries was observed, however, at any age (Fig. 4BGob). Follicle counts on histological sections were carried out in the population of atretic antral follicles (Fig. 4CGo). Their number was low at 15 dpn and increased thereafter. A nonsignificant decrease in the number of atretic antral follicles was observed on the day of puberty in both groups compared with 21, 28, and 32 dpn. No significant difference was observed between control and irradiated ovaries at any age.



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  		FIG. 4. Comparison of follicular atresia between control and irradiated ovaries. A, Study of follicular atresia by in situ end-labeling in control (Aa) and irradiated (Ab) ovaries at 21 dpn. As shown by the numerous positively labeled granulosa cells in both groups, most of the antral follicles present at that time are atretic (*). Scale bar, 250 µm. B, Comparison of DNA fragmentation from 21 dpn to first estrus between control and irradiated ovaries. Ba, Autoradiogram of a representative Southern blot. C, Control; I, irradiated; L, DNA molecular weight ladder. DNA was 3' end-labeled with digoxigenin-dideoxy-UTP, and 1 µg was electrophoresed on 2% agarose gel as described in Materials and Methods. Bb, Quantitation of low molecular weight DNA (fragments <1.2 kb) in control ({blacksquare}) and irradiated ({square}) ovaries. Bars represent the mean ± SEM of three independent experiments conducted in triplicate. Data were analyzed by two-way ANOVA. No significant differences were observed between the two groups at any age. Effect of the treatment, P < 0.0001; effect of age, P < 0.0001; treatment by age, P = 0.06. Within each group, different letters indicate significant differences between ages. C, Number of atretic antral follicles in control ({blacksquare}) and irradiated ({square}) ovaries. Follicle counts were performed on histological sections as described in Materials and Methods. Bars represent the mean ± SEM of three ovaries at 15 dpn and four ovaries at the other ages. Data were analyzed by two-way ANOVA. No significant differences were observed between the two groups at any age. Effect of the treatment, P = 0.03; effect of age, P < 0.0001; treatment by age, P = 0.34. Within a group, different letters indicate significant differences between ages.

 
Follicular maturation
Several markers known to be involved in follicular maturation, namely inhibin {alpha}- and ßA-subunits (23), P450 aromatase, which converts androgens to estrogens (9), and LH receptor (24, 25, 26), were studied throughout the immature period by in situ hybridization. The observations, summarized in Table 4Go and illustrated at 15 dpn and first proestrus in Fig. 5Go, indicated that there were no differences in the expression patterns of all four genes between irradiated and control females.


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  		TABLE 4. Expression pattern of follicular maturation markers throughout the immature period in control and irradiated ovaries by in situ hybridization

 


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  		FIG. 5. In situ analysis of follicular maturation in control and irradiated ovaries at 15 dpn and first proestrus. In situ hybridization for inhibin {alpha}-subunit (inh {alpha}; A–D), inhibin ßA-subunit (inh ßA; E–H), P450 aromatase (P450-Aro; I–L), and LH receptor (LH-R; M–P). At 15 dpn, inh {alpha} (A and B) inh ßA (E and F), and P450-Aro (I and J) are expressed in primary (black arrow), preantral (black arrowhead), and antral (white arrowhead) follicles. Atretic follicles (*) stain for inh {alpha} and display no or weak staining for inh ßA and P450-Aro. Inh {alpha} (A and B) and LH-R (M and N) are expressed in thecal cells and interstitial glands (white arrows). At first proestrus, inh {alpha} (C and D) is widely expressed by the different categories of growing follicles. Inh ßA (G and H) and P450-Aro (K and L) are observed in PoF, but not in preantral (black arrowheads), antral (white arrowhead), or atretic follicles (*). LH-R (O and P) is found in the granulosa cell layers of PoF and in thecal cells and interstitial glands (white arrow). Scale bar, 150 µm.

 
In both groups, inhibin {alpha}-subunit was expressed in the granulosa cell layers of all follicles from the small primary follicle stage and, to a lesser extent, in thecal and mesenchymal cells (Fig. 5Go, A–D). In large atretic antral follicles, its expression became restricted to perioocyte cells. At first proestrus, staining for inhibin {alpha}-subunit confirmed that very few preantral and antral follicles remained in the irradiated ovaries. Preovulatory follicles (PoF) were present at this time in both groups (Fig. 5Go, C and D). This is in agreement with the observation of fresh corpora lutea at first estrus. Inhibin ßA-subunit (Fig. 5Go, E–H) and P450 aromatase (Fig. 5Go, I–L) were expressed in an overlapping pattern throughout the immature period in both groups. At 5 dpn, they were expressed in the granulosa cell layer of growing follicles from the primary stage, and their staining progressively increased in both preantral and antral follicles in infantile ovaries in the two groups. At 21 dpn and in older animals, inhibin ßA-subunit and P450 aromatase were no longer expressed by all growing follicles, but were confined to healthy large antral follicles (Table 4Go). At first proestrus, their expression was restricted to PoF in both groups (Fig. 5Go, G, H, K, and L).

LH receptor expression was detected from 8 dpn in the thecal layer of preantral and antral follicles and in sparse cells in the mesenchymal compartment (Fig. 5Go, M–P). From 21 dpn, this gene was highly expressed in the well-developed interstitial glands, which were mainly distributed in the center of the ovary in both groups (Fig. 5Go, O and P). In addition, LH receptor was expressed in the granulosa cell layers of PoF at first proestrus (Fig. 5Go, O and P).

Plasma levels of E2, T, and inhibin B
To determine whether the fall in the number of growing follicles altered the functional maturation of the ovary, dosages of plasma E2, T, and inhibin B were performed from 21 dpn up to first estrus (Fig. 6Go, A–C). In both groups, concentrations of plasma E2 were constant between 21 and 32 dpn, but showed a high variability at this latter age (Fig. 6AGo). On the day of puberty, the levels of circulating E2 showed a nonsignificant decrease compared with those of 32 dpn females. Levels of plasma T were constant throughout the period studied (Fig. 6BGo). In both control and irradiated animals, concentrations of plasma inhibin B decreased steeply between 21 and 28 dpn, and females displayed low levels until 32 dpn. High levels were found in both groups at first estrus (Fig. 6CGo). For all three hormones, no significant differences were observed between irradiated and control females.



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  		FIG. 6. Plasma concentrations of E2 (A), T (B), and inhibin B (C) in control ({blacksquare}) and irradiated ({square}) females from 21 dpn to first estrus. Bars represent the mean ± SEM. Numbers in brackets indicate the number of plasma pools (two or three animals per pool) at 21 and 28 dpn or of individual samples at 32 dpn and first estrus for both E2 and T, and of individual samples at every age for inhibin B. Data were analyzed by two-way ANOVA. No significant differences were observed between the two groups at any age. A: Effect of the treatment, P = 0.7; effect of age, P = 0.02; treatment by age, P = 0.98. B: Effect of the treatment, P = 0.88; effect of age, P = 0.12; treatment by age, P = 0.89. C: Effect of the treatment, P = 0.04; effect of age, P < 0.0001; treatment by age, P = 0.70. Within a group, different letters indicate significant differences between ages.

 
Premature reproductive senescence
From 2 months of age, irradiated and control females (n = 11 in each group) were continuously mated with males to evaluate their reproductive capacity (Table 5Go). Within the following 2 months, every control female had two litters; the interval between the beginning of the mating and the first litter was 25.1 ± 1.2 d, and for the second litter it was 23.1 ± 0.35 d. For 72.3% (8 of 11) of the irradiated females, the interval between the beginning of the mating and first litter was increased compared with that in controls, reaching 31.4 ± 2.3 d. One irradiated female had its first litter 3 months after the beginning of the mating, and 2 others never conceived. Moreover, within the 2 months, only 1 of 11 irradiated females had 2 litters. This impaired reproductive potential was also apparent in the reduction of the litter size (3.4 ± 0.6 vs. 10.7 ± 1.4 pups for controls in the first litter). Irradiated females conceived pups with no morphological abnormality. As in control females, the number of embryonic implantation sites in the uterus was in accordance with the number of newborn pups (data not shown). Both macroscopic and microscopic observations of the irradiated ovaries at 2 months showed corpora lutea and very few growing follicles in contrast to those of controls (Fig. 7Go, A and B). Between 4–6 months, irradiated ovaries were small compared with control ovaries, and a few of them contained hemorrhagic and follicular cysts (Table 5Go and Fig. 7Go, C and D). At this age, they contained essentially interstitial cells, fibrous tissue, and old corpora lutea (Fig. 7DGo). At 6 months, ovarian and uterine weights were reduced by 60.4% and 21%, respectively, and pituitary weights were increased by 17.4% in the irradiated females compared with the controls (Table 6Go).


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  		TABLE 5. Reproductive performance of control and irradiated females in continuous mating

 


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  		FIG. 7. Ovarian senescence. Histological sections of control (A) and irradiated (B) ovaries at 2 months. Note the few growing follicles (black arrows) in irradiated ovaries compared with controls. C, Gross morphology of control (left) and irradiated (right) ovaries at 6 months. Note the presence of hemorrhagic (white arrows) and follicular cysts (*) in irradiated ovary. D, Histological sections of irradiated ovaries at 6 months. This ovary is devoid of follicles and enlarged by a follicular cyst (*). Scale bar, 500 µm.

 

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  		TABLE 6. Weights of ovary, pituitary, and uterus in control and irradiated females at 6 months

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Our results, in agreement with previous studies, show that oocytes contained in primordial follicles are particularly sensitive to ionizing radiation, whereas growing oocytes are relatively radioresistant (13, 14). We have demonstrated that whole body irradiation performed at 5 dpn with a single dose of 1.5 Gray induced a near complete deletion of the primordial follicle pool within a time period of 24 h. Indeed, at 6 dpn, the irradiated ovary was endowed with less than 1% of healthy primordial follicles compared with controls. The treatment was also toxic for part of the primary follicles, in particular the smaller ones, as shown by the increased oocyte attrition observed 24 h after irradiation. In contrast, the other growing follicles present at 5 dpn, namely large primary and preantral follicles, remained unaffected. At 6 dpn, their number was similar to that of controls. The radiotolerance of these growing follicles was further supported by the fact that no increased cellular apoptosis was found in the irradiated ovary 48 h after treatment. In addition to their sensitivity to irradiation, oocytes contained in primordial follicles are particularly susceptible to environmental chemicals and chemotherapeutic drugs (27, 28). Several genes have been involved in the process of oocyte attrition, such as members of the Bcl-2 family (5, 29, 30, 31, 32). The sphingomyelin pathway has recently been demonstrated to be of major importance in both physiological and irradiation-induced death of the female germ cells (33). The reason for the radiosensitivity of quiescent oocytes and, conversely, the relative resistance of growing oocytes is unknown. Further investigations will have to establish the relationship between the activation of the oocyte death program after irradiation and the follicular stage.

Although the onset of puberty took place similarly in control and irradiated females, the reproductive capacity of irradiated females was severely impaired from the beginning of reproductive life. At 2 months of age, females displayed a severe reduction in their litter size, and by 4 months, most of them had become sterile despite continuous mating. Even if the influence of conception defects due to irradiation damage to oocytes cannot be excluded, it is noticeable that premature reproductive senescence may be closely related to the dramatic reduction of follicular content, as evidenced by the sterile aspect of adult ovaries. Our study of follicular development and dynamics revealed that ovarian deficiency was already becoming established as early as the immature period. This was illustrated by a sequential fall in the number of growing follicles in each category, first obvious for primary follicles at 15 dpn, for preantral follicles at 28 dpn, and for antral follicles at 32 dpn. On the day of puberty, the pool of preantral follicles was nearly exhausted, and the number of antral follicles was reduced by more than 80%. Primary and preantral follicles surviving irradiation, however, developed with a growth pattern similar to that of follicles in control ovaries. As shown by PCNA staining, numerous granulosa cells in primary and preantral follicles were proliferating at 6 dpn. In both groups, moreover, the first antral follicles were observed at 8 dpn. The assumption that the proliferation of follicular cells was unaltered was further supported by the fact that a similar number of preantral and antral follicles was found in both groups until 21 and 28 dpn respectively, i.e. until the nonrenewal of the growing follicle pool became obvious.

Our complementary analysis of the functional maturation of follicles by in situ hybridization confirmed that the development of growing follicles was similar in irradiated and control ovaries throughout the immature period. Granulosa cells displayed the same spatio-temporal expression pattern of genes encoding P450 aromatase and inhibin {alpha}- and ßA-subunits, while from 8 dpn, as already reported (25), thecal cells characteristically expressed the LH receptor gene. In agreement with previous studies of normal ovarian development (9, 34), between 15 and 21 dpn we observed changes in the expression pattern of P450 aromatase and inhibin ßA-subunit genes in both irradiated and control ovaries. From 5–15 dpn, i.e. during neonatal and infantile periods (35), these two genes were expressed by all growing follicles from the primary follicle stage on. In contrast, at 21 dpn, i.e. the beginning of the juvenile period (35), and thereafter, their expression was restricted to healthy large antral follicles. As animals reached puberty, only preovulatory follicles expressed both genes. From our observations in irradiated ovaries it may be concluded that the absence of staining for the transcripts for P450 aromatase and inhibin ßA-subunit in healthy preantral follicles from the juvenile period on results from down-regulation. Indeed, as no recruitment of new follicular waves took place in the irradiated ovaries, the preantral follicles negatively stained at 21 dpn undoubtedly originated from the pool of preantral follicles positively stained at 15 dpn. During the transition between infantile and juvenile periods, major changes are known to occur in endocrine balance as well as in follicular dynamics. Thus, the levels of circulating FSH, initially high during the infantile period, fall during the juvenile period (36, 37) under the control of inhibin and estradiol negative feedback (38, 39). Concomitantly, the follicular atresia process begins, leading to the elimination of numerous growing follicles and, in particular, of follicles at the antral stage (4). Taking into account that as early as during the neonatal period the expression of P450 aromatase (40), and inhibin ß-subunits (41) genes is gonadotropin dependent, it can be assumed that the variations in the levels of FSH are responsible for the changes in the expression of P450 aromatase and inhibin ßA-subunit genes.

The shift in gene expression pattern occurring similarly in irradiated and control ovaries indicates that the regulative event(s) related to the transition between infantile and juvenile periods has taken place normally in irradiated females. Completion of ovarian maturation was evidenced, moreover, by the unaltered levels of E2, T, and inhibin B and the normal onset of puberty in terms of age and number of ovulated follicles at first estrus. Thus, at puberty the irradiated ovary seems to adhere to the law of follicular constancy (42), according to which the ovulation number is constant in a given species, even when the ovarian mass, and thus the number of follicles, are experimentally reduced. In the case of unilaterally ovariectomized (ULO) rats, compensatory ovulation in the remaining ovary results from changes in follicular dynamics taking place in response to modifications of endocrine balance. For example, in late prepubertal ULO, full compensation of ovulation number was achieved at first estrus by decreased follicular atresia and at second estrus by increased recruitment of antral follicles (43). In our analysis of follicular atresia examining in situ the apoptotic process by TUNEL assay, no significant difference between irradiated and control ovaries was observed. In both groups the first atretic follicles appeared at 12 dpn, and the evolution of follicular atresia proceeded similarly at all ages. This was further confirmed by quantitative studies performed by follicular counts on histological sections and Southern blot analysis of DNA fragmentation. In irradiated ovaries, despite the progressive decrease in the number of healthy growing follicles, no parallel reduction in the number of atretic follicles was observed. In other words, there was no census mechanism detecting the deficit in growing follicles and limiting the process of follicular atresia.

Together, our results show that the development of initial follicular waves proceeded similarly as that of their counterparts in controls regarding follicular growth and atresia and proved sufficient to trigger the first ovulation. In prepubertal normal rats, the selection of antral follicles destined to ovulate is considered to take place 7–8 d before puberty (44). In irradiated females, at the time of selection (~28 dpn), the pool of antral follicles was quantitatively similar to that in controls and was therefore capable of ensuring the selection of a normal number of ovulatable follicles. The normal recruitment of follicles destined to ovulate was further demonstrated by the normal levels of E2 and inhibin B until puberty.

In contrast with ULO rats (43) and rats with dysgenesic ovaries (8, 9), in neonatal irradiated rats the deficit in follicles induced no modifications in the dynamics of growing follicles. In the two former models, the entire follicular complement was altered by the treatment. In contrast, in neonatal irradiated females, essentially primordial follicles were affected, and the development of the initial follicular waves proceeded normally. Through their differentiation and their functionality, these growing follicles may well account for the endocrine capacity of the maturing ovary.

In conclusion, with our initial purpose to investigate the influence of neonatal deletion of the primordial follicular pool, we developed an experimental model that allowed us to study more closely the differentiation and dynamics of the subset of follicles initiating their growth shortly after birth. Our results provide strong evidence that the initial follicular waves are sufficient to ensure ovarian maturation until puberty and thus contribute to the onset of reproductive life.


   Acknowledgments
 
We greatly thank O. Locquet for technical assistance, and M. P. Monneret for help with the steroid RIA. We are grateful to Drs. H. Coffigny and R. Counis for valuable discussion, Dr. J. P. Lefaix for animal irradiation, and S. Bouret for helpful assistance with statistical analyses.


   Footnotes
 
This work was supported by Electricité de France and the Ministère de l’Education Nationale et de la Recherche Scientifique et Technique, France. C.J.G. is the recipient of a fellowship from the Ministère de l’Education Nationale et de la Recherche Scientifique et Technique.

Abbreviations: dpn, Days postnatally; E2, 17ß-estradiol; Gy, Gray; PCNA, proliferating cell nuclear antigen; PoF, preovulatory follicles; T, testosterone; SSC, sodium saline citrate; T, testosterone; TUNEL, terminal deoxynucleotidyltransferase-mediated deoxy-UTP-fluorescein nick end labeling; ULO, unilaterally ovariectomized.

Received January 16, 2003.

Accepted for publication April 17, 2003.


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Top
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 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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