Biol Reprod Track the topics, authors and articles important to you
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Abdennebi, L.
Right arrow Articles by Monniaux, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Abdennebi, L.
Right arrow Articles by Monniaux, D.
Agricola
Right arrow Articles by Abdennebi, L.
Right arrow Articles by Monniaux, D.
Biology of Reproduction 60, 845-854 (1999)
©Copyright 1999 Society for the Study of Reproduction, Inc.

Comparative Expression of Luteinizing Hormone and Follicle-Stimulating Hormone Receptors in Ovarian Follicles from High and Low Prolific Sheep Breeds1

L. Abdennebia, P. Mongetb, C. Pisseletb, J.J. Remya, R. Salessea, and D. Monniaux2,b

a Unité Récepteurs et Communication Cellulaire I.N.R.A. Biotechnologies, 78352 Jouy-en-Josas, France b Station I.N.R.A. de Physiologie de la Reproduction des Mammifères Domestiques, 37380 Nouzilly, France


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Expression of gonadotropin receptors and granulosa cell sensitivity to gonadotropin hormones by small (1–3 mm) and large (3.5–7 mm) follicles were compared in Romanov (ROM, ovulation rate = 3) and Ile-de-France (IF, ovulation rate = 1) ewes in the early and late follicular phase. In healthy follicles, LH receptor levels in granulosa cells increased with increasing follicular size (p < 0.001) while FSH receptor levels decreased (p < 0.05). In granulosa cells of large follicles, LH receptor (LHR) mRNA levels were greater in the late than in the early follicular phase (p < 0.001, p < 0.05, for ROM and IF, respectively). In the early follicular phase, LHR levels in granulosa (p < 0.001) and theca cells (p < 0.05) of small follicles were greater in ROM than in IF ewes. FSH receptor mRNA levels in granulosa cells of small and large ROM follicles were greater than in the corresponding IF follicles (p < 0.05). Finally, a greater responsiveness (increase in cAMP secretion) to both FSH and hCG was observed by granulosa cells collected during the early follicular phase from ROM vs. IF ewes. Data provide evidence that the greater ovulation rate in the ROM as compared to the IF breed is associated with a greater gonadotropin responsiveness during the early follicular phase.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In the ewe, ovulation rate varies between breeds from 1 to 5, with the exception of the merino ewes homozygous for the Booroola gene (ovulation rate greater than 5). Ovulation rate is the most important determinant of litter size [1] and is positively correlated to the number of large estrogenic follicles during the estrous cycle and in periods of anestrus [2, 3]. Ovaries from prolific Romanov (ROM) ewes (ovulation rate 3–5) have more growing antral follicles than do those from the nonprolific Ile-de-France (IF) breed (ovulation rate 1–2) [4], and the differences in ovulation rate between these breeds are associated with differences in the pattern of terminal follicular development [5]. In most studies, no clear differences in plasma concentrations of LH and FSH between breeds with different ovulation rates have been found [6–8] even though slightly higher plasma concentrations of FSH have been reported at some stages of the estrous cycle in prolific breeds [9–11]. It is possible instead that prolific and nonprolific breeds are characterized by differences in sensitivity of follicular cells to gonadotropins. In ovaries, FSH receptors are expressed by granulosa cells [12, 13] and are detected in ewes in the earliest stages of follicular growth [14]. In addition, during terminal follicular development, LH receptors are expressed in theca cells from antral follicles and highly expressed in granulosa cells from antral follicles larger than 3 mm in diameter [15].

No data are available concerning expression and functionality of gonadotropin receptors in prolific and nonprolific breeds. In this study, we tested the hypothesis that differences in receptivity of follicular cells to gonadotropins are related to differences in mechanisms of selection of ovulatory follicle(s). We thus compared prolific ROM and nonprolific IL breeds for 1) expression of gonadotropin receptor mRNAs and proteins in follicular cells in vivo and 2) responsiveness of granulosa cells to gonadotropin hormones in vitro. Comparisons were performed at two stages of the follicular phase, i.e., before and after the selection of ovulatory follicles had occurred [5].


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals and Sample Collection

Twenty-six adult cyclic ROM and 28 IF ewes were treated with progestogen (intravaginal flurogestone acetate sponges, 40 mg; Intervet, Angers, France) to synchronize estrus. Ewes were slaughtered either 12 h (early follicular phase) or 36 h (late follicular phase) after sponge removal, and ovaries were collected within 5 min after slaughter. With this treatment, the LH preovulatory surge occurred 40–44 h after sponge removal. At each stage of the follicular phase, 4 ewes per breed were used for in situ hybridization experiments, and 3 ROM and 4 IF ewes were used for in situ ligand binding experiments. In both experiments, immediately after slaughter, ovaries were collected and immediately embedded in Tissue Tek (Miles Laboratories, Elkhart, IN), frozen in liquid nitrogen, and stored at -80°C until use. For studies of cAMP production by gonadotropin-stimulated granulosa cells in vitro, 12 ewes per breed were used in 6 separate cultures (3 cultures at each stage of the follicular phase). For each culture and each stage of the follicular phase, granulosa cells from 2 ewes per breed were pooled.

Preparation of Sections for In Situ Hybridization, In Situ Binding, and Morphologic Assessment of Follicles

Frozen ovaries were serially sectioned at a thickness of 10 µm with a cryostat, and sections were mounted on microscope slides. For in situ hybridization experiments, sections were fixed with 4% paraformaldehyde in PBS (0.01 M, pH 7.4) containing 15 mM vanadyl ribonucleoside complex (RNase inhibitor; Gibco Life Technologies, Cergy-Pontoise, France) for 10 min at room temperature, washed two times in PBS, and incubated in PBS containing 0.0025% Triton X-100 (v:v) for 15 min at room temperature. The slides were washed three times in PBS; then sections were dehydrated in increasing concentrations of ethanol (30%, 50%, and 70% in 0.3 M ammonium acetate) and stored in 70% ethanol until hybridization. For in situ ligand binding experiments, sections were immediately fixed for 10 min at 4°C in a solution containing 2% formaldehyde and picric acid, washed three times in cold PBS, air-dried, and stored at -20°C until binding experiments.

Histological Determinations of Follicular Size and Quality

Adjacent sections were fixed in methanol-formaldehyde-acetic acid (80:15:15), and subsequently stained with Feulgen (Merck, Schuchardt, Germany). The quality of each follicle was assessed by microscopic examination. Follicles were morphologically classified as healthy or atretic. Follicles were judged healthy when they showed frequent mitosis and no or a few pyknotic bodies in the granulosa cells. Atretic follicles exhibited clear degenerative changes such as the presence of frequent pyknotic bodies and local destruction of basement membrane. Late atretic follicles, in which granulosa cells had almost completely disappeared, were not included in this study. Follicles were classified according to size: small follicles (S, 1–3 mm in diameter) and large follicles (L, 3.5–7 mm in diameter); a follicular size of 3 mm was chosen as a limit of size class because it corresponds to acquisition of LH receptors in sheep granulosa cells [15].

In Situ Hybridization

Complementary RNA probe preparation The porcine LH receptor (LHR) full-length cDNA cloned into the transcription vector pBluescript (Stratagene, La Jolla, CA) and the HindIII-BamHI 380 FSH receptor (FSHR) fragment contained within the extracellular domain [16] and subcloned in pGEM3z(-) vector (Promega, Madison, WI), were used as templates for the synthesis of antisense and sense RNA probes. The linearized plasmid (1 µg) was incubated in the presence of 500 µM ribonucleotide (r) ATP, rGTP, 50 µCi of [35S]CTP, and 50 µCi of [35S]UTP (Dupont de Nemours NEN, les Ulis, France) in the presence of T3 or T7 RNA polymerase (Promega) for 1 h at 37°C. After incubation with DNase (RQ1; Promega) to remove DNA, cRNA probes were purified on a Sephadex G50 (Pharmacia, Uppsala, Sweden) column to separate cRNA from free nucleotides. The cRNA probes were then frozen at -80°C and used for in situ hybridization within 1–2 days.

Hybridization Procedures

In situ hybridization was performed as previously described [17]. Briefly, sections were incubated for 2 h at 50°C in a hybridization buffer containing 50% formamide (Merck, Nogent-sur-marne, France), 0.6 M NaCl, 10 mM Tris, 1 mM EDTA, 1% SDS (Serva BioWhittaker, Fontenay-sous-Bois, France), 10 mM dithiothreitol (DTT; Boehringer Mannheim, Meylan, France), 250 µg/ml tRNA (Sigma, l'Isle-d'Abeau-Chesnes, France), 2% Denhardt's reagent (Eurogentec, Angers, France), and 100 mg/ml polyethyleneglycol 6000 (Prolabo, Fontenay-sous-Bois, France). Hybridization was performed by covering sections with hybridization buffer containing 200 000 cpm labeled probes at 50°C overnight. After hybridization, slides were incubated successively in PBS containing 5 mM MgCl2 for 5 min at room temperature and in Tris buffer (10 mM Tris, 0.5 M NaCl, pH = 8) containing 20 µg/ml RNase A (Boehringer Mannheim), two times for 30 min at 37°C. Then slides were washed successively in 1) Tris buffer without RNase A for 30 min at 37°C; 2) in 50% formamide, 1 mM DTT, and double-strength SSC (single-strength SSC is 150 mM sodium chloride, 15 mM sodium citrate, pH 7) for 30 min at 50°C; 3) in 50% formamide, 1 mM DTT, and single-strength SSC for 30 min at 50°C; 4) in 50% formamide, 1 mM DTT, single-strength SSC, and 0.05% Triton X-100 for 30 min at 37°C; and 5) in 50% formamide, 1 mM DTT, 0.1-strength SSC, and 0.05% Triton X-100 for 30 min at 37°C. Sections were then dehydrated in ethanol (30%, 50%, and 70% in 0.3 M ammonium acetate), air-dried, dipped in autoradiographic K5 emulsion (IIford, St. Priest, France), and exposed for 30 days at 4°C in a dark box. After the autoradiographs were developed, the sections were counterstained with hematoxylin and mounted for microscopic examination and quantitative analysis of labeling.

In Situ Ligand Binding

In situ ligand binding was performed as previously described [18, 19]. Sections were incubated for 5 h at room temperature in drops of PBS (0.1% BSA, pH 7) containing 400 000 cpm/100 µl of either 125I-hFSH (Dupont de Nemours NEN; specific activity 90–200 µCi/µg) or 125I-hCG (NIH, batch CR127, specific activity 29 µCi/µg). In adjacent sections, nonspecific binding for each ligand was obtained after incubation of labeled hormones together with an excess of unlabeled FSH or hCG (both 800 ng/100 µl). After incubation, sections were washed twice in PBS for 5 min at 4°C, air-dried, fixed in 3% glutaraldehyde-PBS at 4°C for 15 min, washed again four times for 5 min in PBS at 4°C, and air-dried. Slides were dipped in autoradiographic emulsion (Kodak NTB-2 liquid emulsion [Eastman Kodak, Rochester, NY], diluted 1:1 in distilled water) and exposed for 2 wk at 4°C. After development of the autoradiographs, sections were counterstained with hematoxylin and mounted for microscopic examination and quantitative analysis of labeling.

Quantitative Analysis of Labeling

For both in situ hybridization and in situ ligand binding, quantitative analysis of labeling was performed as previously described [17], using a microscope-linked PC-based image analyzer (Visilog 4.1.5; Noesis, Velizy, France). Each section was analyzed with a x100 objective. Labeling was quantified by measuring the area occupied by silver grains present in a constant area (45 µm2) of tissue (granulosa, G, or theca, T) section of each follicle. Labeling was estimated from 40 measurements randomly distributed on four parts of a section for each follicular compartment (granulosa or theca) of each analyzed follicle. In granulosa cells, quantification was performed close to the basal membrane. For in situ hybridization experiments, data for hybridization intensity were obtained by subtracting values measured on the section hybridized with the sense probe from values measured on the section hybridized with the antisense probe on equivalent areas of adjacent sections. For in situ binding experiments, specific labeling was obtained by subtracting the nonspecific binding values from the total number of grains on equivalent areas of adjacent sections.

In Vitro cAMP Production

After slaughter, ovaries were placed in Medium B2 [20] and transported to the laboratory. Small (1–3 mm) and large (3.5–7 mm) individual follicles were dissected and measured with a millimeter scale. For each follicle, the antral fluid was removed, and the collapsed follicle was placed open in 100 µl of medium B2. Granulosa cells were removed by scraping the interior surface of the follicle wall with a platinum loop, and were pooled in a glass tube according to follicle size (small or large). Both cell suspensions were washed by centrifugation (300 x g, 7 min) and resuspended in 1 ml of medium B2. An aliquot was removed to determine cell number using a hemocytometer and cell viability after vital staining with trypan blue dye (0.125%, final concentration). Aliquots of 50 000 viable cells from small or large follicles were incubated for 2 h (final volume = 500 µl) with increasing concentrations (0.5, 1, 5, 10, 50, 100 ng/ml) of hCG (Chorulon, kind gift from Intervet) or recombinant human FSH (Gonal-Fr; Serono, Boulogne, France) in the presence of 0.1 mM of the phosphodiesterase inhibitor 1-methyl-3-isobutylxanthine (Sigma). For each hormonal concentration, triplicate incubations were performed. After centrifugation at 300 x g for 7 min, the supernatants were collected and stored at -20°C until cAMP assay. Cyclic AMP levels were measured in supernatants by RIA (Dupont de Nemours NEN) after extraction with ethanol according to a published procedure [21]. The intra- and interassay coefficients of variation for the cAMP assay were less than 10%.

Statistical Analysis

For in situ hybridization and ligand binding experiments, statistical analysis of the results was performed using ANOVA. Different models were used to determine the effect of follicular size, stage of the follicular phase, and breed on the expression of LHR and FSHR mRNA and on ligand binding. Duncan's post-hoc multiple-range test was used to compare means. In analyzing the effects of atresia on LHR and FSHR mRNA and on ligand binding, data from all healthy and atretic follicles, irrespective of breed and stage of follicular phase, were considered. To test the effects of follicular size, breed, and stage of the follicular phase on expression of LHR and FSHR mRNA and on ligand binding, only data from healthy follicles were considered. Data from the two breeds and the two stages of follicular phase were pooled to test the effect of size on FSH receptor levels. In the studies of cAMP production, we tested the effect of hormone concentration, breed, and stage of the follicular phase for each class of follicle. Differences in sensitivity to gonadotropins were tested by comparing cAMP values obtained for each hormone concentration with values obtained in basal levels, by using Student's t-test. In addition, responsiveness to each gonadotropin was assessed by calculating the slope (b) of the regression line between the logarithm of the dose of gonadotropin and the concentration of cAMP in the incubation medium. Slopes were compared by t-test. Proportions of atretic and healthy follicles in each class of size and breed were compared by chi-square test. All results are reported as mean ± SEM. Differences with p > 0.05 were considered as nonsignificant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Characteristics of Follicular Populations

A total of 233 follicles from both breeds were histologically characterized for their quality (atretic or healthy) and size (small or large). At 12 h after sponge removal, there was no difference between breeds in the proportion of atretic follicles when small follicles were considered (64%, n = 61, atretic IF follicles vs. 55%, n = 69, atretic ROM follicles). However, at that time, the proportion of atretic large follicles was greater among ROM ewes than among IF ewes (40%, n = 20, vs. 10%, n = 10, respectively, p < 0.05). At 36 h after sponge removal, no significant differences between breeds were observed for proportions of atretic follicles in small (44%, n = 17, IF vs. 51%, n = 33, ROM) or in large follicles (40%, n = 10, IF vs. 31%, n = 13, ROM).

Expression of mRNAs for LH and FSH Receptors

LH receptor mRNAs Irrespective of the stage of follicular development and breed, in small antral follicles, LHR mRNAs were clearly present in theca interna cells but low levels were also detected in granulosa cells (Fig. 1). In large antral follicles, LHR mRNA levels were highly expressed in both granulosa and theca interna cells (Fig. 1). In small follicles, LHR mRNA levels were less abundant in theca cells from atretic follicles as compared to healthy follicles (235 ± 19, n = 58, atretic follicles vs. 361 ± 21, n = 47, healthy follicles; p < 0.001). In large follicles, LHR mRNA expression was less abundant in both granulosa and theca cells of atretic follicles as compared to healthy follicles (216 ± 52, n = 9, atretic granulosa vs. 343 ± 47, n = 17, healthy granulosa, p < 0.05; and 302 ± 59, n = 9, atretic theca vs. 596 ± 44, n = 17, healthy theca, p < 0.001). In healthy follicles (irrespective of the breed of sheep and stage of the follicular phase), LHR mRNAs increased both in granulosa and theca cells with increasing follicular size (130 ± 17, n = 47, small granulosa vs. 361 ± 24, n = 17, large granulosa, and 350 ± 29, n = 47, small theca vs. 596 ± 41, n = 17, large theca; p < 0.001).



View larger version (89K):
[in this window]
[in a new window]
  		FIG. 1. In situ localization of LH receptor mRNA (A, F) and FSH receptor mRNA (G, I) in cryosections of follicles from IF (A, C) and ROM (D, I) ewes in the late follicular phase. A, B) Brightfield and darkfield view of a section from a small healthy follicle hybridized with an antisense porcine 35S-labeled LH cRNA probe, showing a higher expression of LHR mRNA in theca cells. C) Darkfield view of the same follicle hybridized with sense 35S-LH cRNA probe. D, E) Brightfield and darkfield view of a section from a large healthy follicle hybridized with antisense porcine 35S-labeled LH cRNA probe, showing a higher expression of LHR mRNA in granulosa and theca cells. F) Darkfield view of the same follicle hybridized with sense 35S-LH cRNA probe. G, H) Brightfield and darkfield view of a section from a small healthy follicle hybridized with antisense porcine 35S-labeled FSH cRNA probe showing a higher expression of FSH mRNA in granulosa cells. I) Darkfield view of the same follicles hybridized with sense 35S-FSH cRNA probe. G, Granulosa cells; T, theca cells; A, antrum. Scale bar = 50 µm.

The effect of stage of the follicular phase on LHR mRNA expression varied between breeds (Fig. 2, a and b). In ROM ewes, LHR mRNA levels were higher in granulosa (p < 0.001) and theca interna cells (p < 0.01) of large follicles and slightly lower in theca cells of small follicles (p < 0.05) at 36 h than at 12 h after sponge removal. In IF ewes, LHR mRNA levels were higher (p < 0.05) in granulosa cells of large follicles at 36 h than at 12 h after sponge removal. At 36 h after sponge removal, LHR mRNA levels in granulosa cells from large follicles were higher in ROM than in IF ewes (p < 0.05, Fig. 2a). However, at 12 h after sponge removal, no difference between breeds was observed for LHR mRNA expression.



View larger version (25K):
[in this window]
[in a new window]
  		FIG. 2. LH receptor mRNA levels in granulosa and theca of healthy follicles from ROM and IF ewes. The level of expression, in arbitrary units (mean ± SEM), was determined in granulosa (a) and theca cells (b) of small and large healthy follicles, recovered 12 h (early follicular phase) and 36 h (late follicular phase) after progestogen sponge removal. The number of follicles per breed in each class of size and stage of the follicular phase is indicated.

FSH receptor mRNAs FSHR mRNAs were observed only in granulosa and cumulus cells (Fig. 1). FSHR mRNA levels were extremely low compared with LHR mRNA levels. Irrespective of follicular size, breed, or stage of follicular phase, FSHR mRNA levels were lower in atretic follicles than in healthy follicles (34 ± 8, n = 34, vs. 63 ± 7, n = 46, respectively; p < 0.001). In healthy ROM follicles, FSHR mRNA levels were similar in small and large follicles at 12 h and 36 h after sponge removal. However, in healthy IF follicles at 36 h after sponge removal, FSHR mRNA levels were lower in large than in small follicles (p < 0.05, Fig. 3). In addition, in small healthy IF follicles, FSHR mRNA levels were higher at 36 h than at 12 h after sponge removal (p < 0.01). Finally, when breeds were compared, FSHR mRNA levels were higher in small and large ROM follicles than in small and large IF follicles at 12 h (p < 0.02 and p < 0.05, respectively), but not at 36 h after sponge removal.



View larger version (33K):
[in this window]
[in a new window]
  		FIG. 3. FSH receptor mRNA expression in granulosa of healthy follicles from ROM and IF ewes. The level of expression, in arbitrary units (mean ± SEM), was determined in granulosa cells of small and large healthy follicles recovered 12 h (early follicular phase) and 36 h (late follicular phase) after progestogen sponge removal. The number of follicles per breed in each class of size and stage of the follicular phase is indicated.

Specific Binding for LH and FSH Receptors

LH/hCG binding sites LH/hCG receptors were detected in theca cells of small and large follicles, and in granulosa cells of large follicles (Fig. 4). Moreover, lower levels were also present in granulosa cells from small follicles. In small follicles, LH/hCG receptor levels were lower in granulosa and theca cells from atretic as compared to healthy follicles (221 ± 24, n = 36, atretic granulosa vs. 311 ± 25, n = 34, healthy granulosa, p < 0.05; and 686 ± 47, n = 36, atretic theca vs. 968 ± 49, n = 34, healthy theca, p < 0.001). In large follicles as well, LH/hCG receptor levels were lower in granulosa and theca cells from atretic as compared to healthy follicles (458 ± 73, n = 6, atretic granulosa vs. 869 ± 40, n =20, healthy granulosa, p < 0.001; and 688 ± 88, n = 6, atretic theca vs. 1018 ± 48, n = 20, healthy theca, p < 0.001). In healthy follicles, LH/hCG receptor levels were greater in granulosa cells but not theca cells from large as compared to small follicles (869 ± 34, n = 20 large granulosa vs. 311 ± 26, n = 34, small granulosa, respectively, p < 0.001). For small healthy IF follicles only, LH/hCG receptor levels in granulosa and theca cells were higher at 36 h than at 12 h after sponge removal (both p < 0.01, Fig. 5, a and b). Finally, at 12 h after sponge removal, LH/hCG receptor levels were greater in granulosa (p < 0.001) and theca interna cells (p < 0.05) from small ROM as compared to small IF follicles, and in theca cells (p < 0.05) from large ROM, as compared to large IF follicles (Fig. 5, a and b).



View larger version (84K):
[in this window]
[in a new window]
  		FIG. 4. Autoradiographic localization of binding sites for LH (A, F) and FSH (G, I) in cryosections of follicles from IF (G, I) and ROM (A, F) ewes in the early follicular phase. A, B) Brightfield and darkfield view of a section from a small healthy follicle incubated in the presence of 125I-hCG. C) Adjacent serial section treated identically except for the addition of an excess of unlabeled hCG. Binding sites were predominantly localized to theca cells (T). D, E) Brightfield and darkfield view of a section from a large healthy follicle incubated in the presence of 125I-hCG. F) Adjacent serial section treated identically except for the addition of an excess of unlabeled hCG. Intense binding was present in granulosa (G) and theca cells (T). G, H) Brightfield and darkfield view of a section from a large healthy follicle incubated in the presence of 125I-FSH. I) Adjacent serial section treated identically except for the addition of an excess of unlabeled FSH. Binding sites were localized to granulosa cells (G). G, Granulosa cells; T, theca cells; A, antrum. Scale bar = 50 µm.



View larger version (25K):
[in this window]
[in a new window]
  		FIG. 5. Binding sites for LH/hCG in granulosa and theca of healthy follicles from ROM and IF ewes. The specific binding for LH/hCG, expressed in arbitrary units (mean ± SEM), was determined in granulosa (a) and theca cells (b) of small and large healthy follicles recovered 12 h (early follicular phase) and 36 h (late follicular phase) after progestogen sponge removal. The number of follicles per breed in each class of size and stage of the follicular phase is indicated.

FSH binding sites FSH receptors were detected only in granulosa cells from small and large follicles (Fig. 4). When all small and large follicles were pooled irrespective of breed of ewe and stage of the follicular phase, FSH receptor levels were lower in large healthy follicles than in small healthy follicles (362 ± 44, n = 19, vs. 503 ± 32, n = 33, respectively; p < 0.01). FSH receptor levels were lower in granulosa cells of small atretic follicles than in small healthy follicles (316 ± 29, n = 37, vs. 503 ± 31, n = 33, respectively; p < 0.001). By contrast to LHR, neither the stage of the follicular phase nor the breed influenced specific FSH binding when all healthy follicles were considered (Fig. 6).



View larger version (30K):
[in this window]
[in a new window]
  		FIG. 6. Binding sites for FSH in granulosa of healthy follicles from ROM and IF ewes. The specific binding for FSH, expressed in arbitrary units (mean ± SEM), was determined in granulosa cells of small and large healthy follicles recovered 12 h (early follicular phase) and 36 h (late follicular phase) after progestogen sponge removal. The number of follicles per breed in each class of size and stage of the follicular phase is indicated.

In Vitro cAMP Production

When ovaries were recovered 12 h after sponge removal, there was no difference between breeds in the cell viability estimates in small and large follicles for the three separate cultures (58% vs. 63% for small ROM vs. small IF, respectively, and 63% vs. 61% for large ROM vs. large IF, respectively). In contrast, at 36 h after sponge removal, cell viability was slightly greater in small and large IF granulosa cells (52% vs. 65% for small ROM vs. small IF, respectively, p < 0.05; and 59% vs. 68% for large ROM vs. large IF, respectively, p < 0.05). At 12 h after sponge removal, granulosa cells from both ROM and IF small follicles showed similar dose-dependent increases in cAMP production when stimulated by FSH (Fig. 7a, slope of the regression line, b = 0.69 ± 0.22 pmol/ng vs. 0.85 ± 0.21 pmol/ng, IF vs. ROM, respectively; p > 0.05), but did not show any response to hCG (Fig. 7b). Similarly, at 36 h after sponge removal, granulosa cells from both small ROM and IF breeds showed similar dose-dependent increases in response to FSH (Fig. 7c, b = 0.85 ± 0.17 pmol/ng vs. 0.99 ± 0.44 pmol/ng, IF vs. ROM, respectively, p > 0.05) but did not respond to hCG (Fig. 7d). In both breeds, granulosa cells from large follicles recovered at 12 h after sponge removal exhibited dose-response increases in cAMP production when stimulated by both gonadotropins (Fig. 8). When granulosa cells from large follicles were recovered at 12 h, responses to FSH (b = 0.62 ± 0.33 pmol/ng vs. 1.69 ± 0.532 pmol/ng, IF vs. ROM, respectively; p < 0.01) and hCG (b = 0.85 ± 0.25 pmol/ng vs. 2.20 ± 0.73 pmol/ng, IF vs. ROM, respectively; p < 0.01) were greater in ROM than in IF ewes (Fig. 8, a and b). Granulosa cells from large ROM and IF follicles recovered at 36 h exhibited a similar dose-response increase in cAMP production in the presence of hCG (b = 1.97 ± 0.18 pmol/ng vs. 1.18 ± 0.3 pmol/ng, IF vs. ROM, respectively; p > 0.05), while FSH stimulated cAMP production of ROM granulosa cells only (b = 0.11 ± 0.21 pmol/ng vs. 0.93 ± 0.14 pmol/ng, IF vs. ROM, respectively, p < 0.01). Finally, in IF ewes at 36 h after sponge removal, response of granulosa cells to FSH was lower in large than in small follicles (0.11 ± 0.21 pmol/ng vs. 0.85 ± 0.17 pmol/ng, respectively, p < 0.01).



View larger version (22K):
[in this window]
[in a new window]
  		FIG. 7. Cyclic AMP production (pmol/50 000 viable cells) by granulosa cells prepared from small follicles of both ROM and IF ewes recovered 12 h (a, b: n = 6 ewes per breed) or 36 h (c, d: n = 6 ewes per breed) after progestogen sponge removal and stimulated by FSH (a, c) or hCG (b, d) for 2 h in vitro. Each datum point is the mean of 3 separate cultures, each culture including triplicate culture wells with cells recovered from 2 ewes. * p < 0.05, ** p < 0.01 compared with nonstimulated cells (basal levels).



View larger version (22K):
[in this window]
[in a new window]
  		FIG. 8. Cyclic AMP production (pmol/50 000 viable cells) by granulosa cells from large follicles of both ROM and IF ewes recovered 12 h (a, b: n = 6 ewes per breed) or 36 h (c, d: n = 6 ewes per breed) after progestogen sponge removal and stimulated by FSH (a, c) or hCG (b, d) for 2 h in vitro. Each datum point is the mean of 3 separate cultures, each culture including triplicate culture wells with cells recovered from 2 ewes. * p < 0.05, ** p < 0.01, *** p < 0.001 compared with nonstimulated cells (basal levels).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we attempted to compare gonadotropin receptor mRNA expression, protein levels, and responsiveness of granulosa cells to gonadotropin hormones, as assessed by increase in cAMP secretion, between low and high ovulating ewes.

Both breeds of ewes used in these studies showed similar changes in gonadotropin receptor expression during follicular growth and atresia. First, atresia was accompanied by a decrease in levels of gonadotropin receptors and their mRNAs in granulosa and theca cells regardless of breed of the ewe and stage of the follicular phase. Similarly, in the sheep and the pig, FSH receptor expression was shown to be reduced in granulosa cells from atretic follicles [14, 22]. Interestingly, LH receptor expression, although reduced, was still observed in atretic follicles of pigs and cattle [22,23]. Second, consistent with previous results in different mammalian species, LH receptor expression was negligible in granulosa cells from small antral follicles and highly enhanced in granulosa cells when follicles increased in size [22–26]. Accordingly, granulosa cells from large but not small follicles produced high levels of cAMP under stimulation by hCG. Third, in theca cells, expression of LH receptors was similar between small and large follicles, in agreement with previous results of LH binding to theca interna tissue in ovine or bovine follicles collected at different stages of the estrous cycle [15, 27, 28]. Finally, FSH receptor levels were shown to be slightly lower in large than in small follicles. Moreover, under FSH stimulation in vitro, granulosa cells from large follicles produced amounts of cAMP lower than (IL ewes) or similar to (ROM ewes) amounts produced by granulosa cells from small follicles. In the pig as well, a negative relationship has been reported between both FSH receptor levels and cAMP production by granulosa cells in vitro, and the size of follicles (between 1 and 6–12 mm) [29, 30]. Conversely, in sheep, Henderson and coworkers [31] showed a 2- to 10-fold increase in cAMP production by granulosa cells stimulated by FSH as the diameter of the follicle increased from 2.5 to 4 mm. The reason for this discrepancy is not known. It could be due to differences between the experiments in the ratio of viable to dead cells.

Some discrepancies were observed between gonadotropin receptor mRNA and protein levels in both breeds. In both ROM and IL ewes, final maturation of large follicles between 12 h and 36 h after sponge removal was accompanied by an increase in LH receptor mRNA but not protein levels. Moreover, at 12 h after sponge removal, the FSHR mRNA level was greater in granulosa cells from small and large follicles of ROM as compared to IL follicles, but the number of FSH binding sites did not differ.

As previously observed, the accumulation of truncated or untranslatable transcripts of gonadotropin receptors [32–34] might explain these differences. In particular, some results [35] suggest that in the case of down-regulation, short RNA transcripts may accumulate but are not translated to protein. Alternatively, the translation machinery of these mRNAs might be prevented.

As stated previously, there was a general agreement between the presence of gonadotropin receptors on granulosa cells and cAMP responsiveness of gonadotropin-stimulated cells. However at 12 h after sponge removal, although the number of FSH and LH binding sites were similar between ROM and IL granulosa cells from large follicles, FSH- and LH-induced cAMP responses were clearly greater by ROM cells. The difference in cAMP production between breeds of ewes could not be attributed to differences in cell viability, since the same number of viable cells (as estimated by trypan blue dye exclusion) were incubated in these in vitro experiments. More likely, these results suggest that the greatest responsiveness of viable granulosa cells in ROM ewes during the early follicular phase might be associated with a more efficient coupling between receptors and adenylate cyclase or to enhanced adenylate cyclase expression levels.

Overall, our results suggest that terminal maturation of follicles occurred earlier in ROM than in IL ewes. First, LH receptor levels in granulosa and theca cells from small follicles were greater in ROM than in Ile-de France ewes at 12 h after sponge removal. Second, at this time, FSH receptor mRNA levels in granulosa cells from small and large ROM follicles were greater than in granulosa cells from the corresponding IL follicles. Finally, at 12 h after sponge removal as well, a greater responsiveness to both FSH and LH of granulosa cells from large ROM follicles, as compared to IL, was observed. Interestingly, Mariana and coworkers [36] have shown that granulosa cells recovered from 6-mo-old ROM lambs were less proliferative and more steroidogenic than cells from IL lambs, after stimulation by FSH in vitro. In Booroola ewes, animals carrying the F gene of prolificacy also acquire aromatase activity and LH receptors at a smaller follicular diameter than do noncarrier animals [37, 38]. In such animals as the ROM breed of ewe, a high ovulation rate seems to be associated with an earlier onset of follicular maturation.

In summary, our results show that the higher ovulation rate among ROM, as compared to IL ewes, is associated with a greater gonadotropin responsiveness of follicles in the early follicular phase, thus allowing these follicles to grow despite a decrease in circulating FSH levels. These data suggest that differences in gonadotropin responsiveness could, at least partly, explain differences in the pattern of terminal follicular development between breeds with different prolificacy.


   ACKNOWLEDGMENTS
 
We thank O. Bastien for his help in the use of the Image Analysis system; A. Solari for statistical assistance; P. Dieudonne for figures; D. Grebert for technical assistance; E. Pajot for helpful comments; and Ismael, Monia, and M. Najar for their support.


   FOOTNOTES
 
1 This work was supported by INRA funding. Back

2 Correspondence: D. Monniaux, Station I.N.R.A. de Physiologie de la Reproduction des Mammifères Domestiques, URA CNRS 1291, 37380 Nouzilly, France. FAX: 3347427743; monniaux{at}tours.inra.fr Back

Accepted: November 10, 1998.

Received: March 19, 1998.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Hanrahan JP, Quirke JF. Contribution of variation in ovulation rate and embryo survival to within breed variation in litter size. In: Land RB, Robinson DW (eds.), Genetics of Reproduction in Sheep. London: Butterworths; 1985: 193–201.
  2. England BG, Webb R, Dahmer MK. Follicular steroidogenesis and gonadotropin binding to ovine follicles during the oestrous cycle. Endocrinology 1981; 109:881–887.[Abstract]
  3. Webb R, Baxter G, McBride D, Ritchie M, Spingbett AJ. Mechanism controlling ovulation rate in ewes in relation to seasonal anoestrus. J Reprod Fertil 1992; 94:143–151.[Abstract]
  4. Cahill LP, Mariana JC, Mauleon P. Total follicle populations in ewes with high and low ovulation rates. J Reprod Fertil 1979; 55:27–36.[Abstract]
  5. Driancourt MA, Gauld IK, Terqui M, Webb R. Variations in patterns of follicle development in prolific breeds of sheep. J Reprod Fertil 1986; 78:565–575.[Abstract]
  6. Findlay JK, Cumming IA. The effect of unilateral ovariectomy on plasma gonadotropin levels, estrus and ovulation rate in sheep. Biol Reprod 1977; 17:178–183.[Abstract]
  7. Adams TE, Quirke JF, Hanrahan JP, Adams BM, Watson JG. Gonadotropin secretion during the preovulatory period in Galway and Finnish Landrace ewes and Finnish Landrace ewes selected for high ovulation rate. J Reprod Fertil 1988; 83:575–584.[Abstract]
  8. Driancourt MA, Philipon P, Locatelli A, Jacques E, Webb R. Are differences in FSH concentrations involved in the control of ovulation rate in Romanov and Ile-de-France ewes. J Reprod Fertil 1988; 83:509–516.[Abstract]
  9. Cahill LP. Folliculogenesis in the sheep as influenced by breed, season and oestrous cycle. J Reprod Fertil 1981; 30:135–142.
  10. Lahlou-Kassi A, Schams D, Glatzel P. Plasma gonadotrophin concentrations during the oestrous cycle and after ovariectomy in two breeds of sheep with low and high fecundity. J Reprod Fertil 1984; 70:165–173.[Abstract]
  11. McNatty KP, Hudson N, Henderson KM, Gibb M, Morrison L, Ball K, Smith P. Differences in gonadotrophin concentrations and pituitary responsiveness to GnRH between Booroola ewes which were homozygous (FF), heterozygous (F+) and non-carriers (++) of a major gene influencing their ovulation rate. J Reprod Fertil 1987; 80:577–588.[Abstract]
  12. Rannikki AS, Zhang FP, Huhtaniemi IT. Ontogeny of follicle-stimulating-hormone receptor gene expression in the rat testis and ovary. Mol Cell Endocrinol 1995; 107:199–208.[CrossRef][Medline]
  13. Richards JS, Ireland JJ, Rao MC, Bernath GA, Midgley AR Jr, Reichert LE Jr. Ovarian follicular development in the rat: hormone receptor regulation by estradiol, follicle stimulating hormone and luteinizing hormone. Endocrinology 1979; 99:1562–1570.[Abstract]
  14. Tisdall DJ, Watanabe K, Hudson NL, Smith P, McNatty KP. FSH receptor gene expression during ovarian follicle development in sheep. J Mol Endocrinol 1995; 15:273–281.[Abstract]
  15. Carson RS, Findlay JK, Burger HG, Trounson AO. Gonadotropin receptors of the ovine ovarian follicle during follicular growth and atresia. Biol Reprod 1979; 21:75–87.[Abstract]
  16. Remy JJ, Mansais YL, Yerle M, Bozon V, Couture L, Pajot E, Grabert D, Salesse R. The porcine follitropin receptor: cDNA cloning, functional expression and chromosal localization of the gene. Gene 1995; 163:257–261.[CrossRef][Medline]
  17. Besnard N, Pisselet C, Monniaux D, Locatelli A, Benne F, Gasser F, Hatey F, Monget P. Expression of messenger ribonucleic acids of insulin-like growth factor binding proteins-2, 4, and 5 in the ovine ovary: localization and changes during growth and atresia of antral follicles. Biol Reprod 1996; 55:1356–1367.[Abstract]
  18. Oxberry BA, Greenwald GS. An autoradiographic study of the binding of 125I-labeled follicle-stimulating hormone, human chorionic gonadotropin and prolactin to the hamster ovary throughout the estrous cycle. Biol Reprod 1982; 27:505–516.[CrossRef][Medline]
  19. Monget P, Monniaux D, Durand P. Localization, characterization, and quantification of insulin-like growth factor-I-binding sites in the ewe ovary. Endocrinology 1989; 125:2486–2493.[Abstract]
  20. Menezo Y. Milieu synthétique pour la survie et la maturation des gamètes et pour la culture de l'oeuf fécondé. C R Acad Sci Paris D 1976; 282:1967–1970.
  21. Morera AM, Saez JM. Mechanisms implicated in ACTH-induced steroidogenic and DNA synthesis refractoriness on adrenal mouse cell line (Y-1). J Steroid Biochem 1980; 12:245–251.[CrossRef][Medline]
  22. Yuan Wei, Lucy MC, Smith MF. Messenger ribonucleic acid for insulin-like growth factors-I and II, insulin-like growth factor-binding protein-2, gonadotropin receptors, and steroidogenic enzymes in porcine follicles. Biol Reprod 1996; 55:1045–1054.[Abstract]
  23. Xu Z, Garverick HA, Smith GW, Smith MF, Hamilton SA, Youngquist RS. Expression of follicle-stimulating hormone and luteinizing hormone receptor messenger ribonucleic acids in bovine follicles during the first follicular wave. Biol Reprod 1995; 53:951–957.[Abstract]
  24. Webb R, England BG. Identification of the ovulatory follicle in the ewe: associated changes in follicular size. Thecal and granulosa cell luteinizing hormone receptors, antral fluids steroids, and circulating hormones during the preovulatory period. Endocrinology 1982; 110:873–881.[Medline]
  25. Peng XR, Hsueh AJW, Lapolt PS, Bjersing L, Ny T. Localization of luteinizing hormone receptor messenger ribonucleic acid expression in ovarian cell types during follicle development and ovulation. Endocrinology 1991; 29:3200–3207.
  26. Bao B, Garverick HA, Smith GW, Smith MF, Salfen BE, Youngquist RS. Changes in messenger ribonucleic acid encoding luteinizing hormone receptor, cytochrome P450-side chain cleavage, and aromatase are associated with recruitment and selection of bovine ovarian follicles. Biol Reprod 1997; 56:1158–1168.[Abstract]
  27. Spicer LJ, Convey EM, Leung K, Short RE, Tucker HA. Anovulation in postpartum suckled beef cows. II. Associations among binding of [125I]-labeled gonadotropins to granulosa and thecal cells, and concentrations of steroids in serum and various sized ovarian follicles. J Anim Sci 1986; 62:742–750.
  28. Henderson KM, Kieboom LE, McNatty KP, Lun S, Heath DA. [125I]-hCG binding to bovine thecal tissue from healthy and atretic antral follicles. Mol Cell Endocrinol 1984; 34:91–98.[CrossRef][Medline]
  29. Lindsay AM, Channing CP. Comparison of the stimulatory effects of ovine, porcine and human follicle-stimulating hormone and of ovine and human luteinizing hormone on the accumulation of cyclic AMP by porcine granulosa cells. J Endocrinol 1979; 80:9–20.[Abstract]
  30. Labarbera AR. Follicle-stimulating hormone (FSH) receptors and FSH-responsive adenosine 3',5'-cyclic monophosphate production in porcine granulosa cells decline with follicular growth. Endocr Res 1994; 20:65–77.[Medline]
  31. Henderson KM, McNatty KP, O'Keefe LE, Lun S, Heath DA, Prisk MD. Differences in gonadotropin-stimulated cyclic AMP production by granulosa cells from Booroola x Merino ewes which are homozygous, heterozygous or non-carriers of a fecundity gene influencing their ovulation rate. J Reprod Fertil 1987; 81:395–402.[Abstract]
  32. Goxe B, Salesse R, Remy JJ, Genty N, Garnier J. LH receptor RNA and protein levels after hormonal treatment of porcine granulosa cells in primary culture. J Mol Endocrinol 1992; 8:119–129.[Abstract]
  33. Koo YB, Ji I, Ji TH. Characterization of different sizes of rat luteinizing hormone/chorionic gonadotropin receptor messenger ribonucleic acids. Endocrinology 1994; 1220:333–337.
  34. Hu Z, Buczko E, Zhuang L, Dufau M. Sequence of the 3' noncoding region of the luteinizing hormone receptor gene and identification of two polyadenylation domains that generate the major mRNA forms. Biochem Biophys Acta 1994; 1220:333–337.[Medline]
  35. Lu DL, Peegel H, Mosier SM, Menon KMJ. Loss of lutropin/human receptor messenger ribonucleic acid during ligand-induced down-regulation occurs post transcriptionally. Endocrinology 1993; 1:235–240.
  36. Mariana JC, Monniaux D, Caraty A, Pisselet C, Fontaine J, Solari A. Immunization of sheep against GnRH early in life: effects on gonadotropins, follicular growth and responsiveness of granulosa cells to FSH and IGF-I in two breeds of sheep with different prolificacy (Romanov and Ile-de-France). Domest Anim Endocrinol 1998; 15:(in press).
  37. Henderson KM, Kieboom LE, McNatty KP, Lun S, Heath D. Gonadotrophin stimulated cyclic AMP production by granulosa cells from Booroola x Romney ewes with and without a fecundity gene. J Reprod Fertil 1985; 75:11–120.
  38. McNatty KP, O'Keeffe LE, Henderson KM, Heath DA, Lun S. [125I]-labelled hCG binding characteristics in theca interna and other tissues from Romney ewes and from Booroola x Romney ewes with and without a major gene influencing their ovulation rate. J Reprod Fertil 1986; 77:477–488.[Abstract]



This article has been cited by other articles:


Home page
 J ANIM SCIHome page
M. C. Lucy
The bovine dominant ovarian follicle
J Anim Sci, March 1, 2007; 85(13_suppl): E89 - E99.
[Abstract] [Full Text] [PDF]

Home page
 Biol. Reprod.Home page
D. Nogueira, C. Albano, T. Adriaenssens, R. Cortvrindt, C. Bourgain, P. Devroey, and J. Smitz
Human Oocytes Reversibly Arrested in Prophase I by Phosphodiesterase Type 3 Inhibitor In Vitro
Biol Reprod, September 1, 2003; 69(3): 1042 - 1052.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
A. Bachelot, P. Monget, P. Imbert-Bollore, K. Coshigano, J. J. Kopchick, P. A. Kelly, and N. Binart
Growth Hormone Is Required for Ovarian Follicular Growth
Endocrinology, October 1, 2002; 143(10): 4104 - 4112.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Endocrinol. Metab.Home page
F. Clement, D. Monniaux, J. Stark, K. Hardy, J. C. Thalabard, S. Franks, and D. Claude
Mathematical model of FSH-induced cAMP production in ovarian follicles
Am J Physiol Endocrinol Metab, July 1, 2001; 281(1): E35 - E53.
[Abstract] [Full Text] [PDF]

Home page
 Endocr. Rev.Home page
E. A. McGee and A. J. W. Hsueh
Initial and Cyclic Recruitment of Ovarian Follicles
Endocr. Rev., April 1, 2000; 21(2): 200 - 214.
[Abstract] [Full Text]

Home page
 Biol. Reprod.Home page
M. Taneja, P. E.J. Bols, A. V. de Velde, J.-C. Ju, D. Schreiber, M. W. Tripp, H. Levine, Y. Echelard, J. Riesen, and X. Yang
Developmental Competence of Juvenile Calf Oocytes In Vitro and In Vivo: Influence of Donor Animal Variation and Repeated Gonadotropin Stimulation
Biol Reprod, January 1, 2000; 62(1): 206 - 213.
[Abstract] [Full Text]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Abdennebi, L.
Right arrow Articles by Monniaux, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Abdennebi, L.
Right arrow Articles by Monniaux, D.
Agricola
Right arrow Articles by Abdennebi, L.
Right arrow Articles by Monniaux, D.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS