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Ontogeny of Follicle-Stimulating Hormone Receptor Gene Expression in Isolated Human Ovarian Follicles¹

Division of Obstetrics and Gynecology, University of Leeds, Leeds General Infirmary, Leeds, West Yorkshire, United Kingdom LS2 9NS

Address all correspondence and requests for reprints to: Kutluk Oktay, M.D., Department of Obstetrics and Gynecology, Division of Reproductive Endocrinology, New York Methodist Hospital, Park Slope, New York 11215. E-mail: koktay{at}netmail.hscbklyn.edu

	Abstract

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FSH stimulates antral follicles to grow, but its role in earlier stages, if any, is obscure. Our aim was to determine the follicle stage at which the FSH receptor (FSHr) gene is first expressed. We used a PCR-based strategy to analyze single follicles ranging from primordial to multilaminar stages after isolation from human ovaries. Ovarian tissue was obtained from 11 women (age range, 25–33 yr) undergoing elective cesarean section. Tissue was partially disaggregated in medium containing 1% collagenase, and follicles were manually dissected free of stroma. Follicle stages were confirmed microscopically as primordial (nongrowing), primary (1 layer of cuboidal granulosa cells), or having 2 or more layers of granulosa cells. Rectus muscle and stromal tissue were used as negative controls. Messenger ribonucleic acid (mRNA) from each follicle was reverse transcribed, and the resulting cDNA was amplified by nested PCR using primers for FSHr and actin. None of the 9 primordial follicles expressed FSHr mRNA. Thirty-three percent of the primary and 2-layer follicles were positive for FSHr mRNA (4 of 12 and 3 of 9, respectively), as were 100% (n = 4) of the multilaminar follicles. The difference in FSH expression between the growing and primordial follicles was significant. This study shows for the first time that transcription of the FSHr gene begins at the earliest stages of follicular growth and indicates that FSH may have a hitherto unsuspected physiological role in preantral follicle development. In addition, this study demonstrates the practical feasibility of investigating the expression of other genes during human folliculogenesis.

	Introduction

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IT IS well established that FSH receptors are present in granulosa cells of growing follicles at preantral and Graafian stages, and that growth and P₄₅₀ aromatase are stimulated by FSH (1). In contrast, the role(s) of FSH at earlier stages of follicular development has not yet been clearly elucidated. There is a general assumption that preantral follicles are not gonadotropin dependent because they are present in animals that are either hypogonadal or whose pituitary glands have been down-regulated with GnRH agonists (2, 3, 4). What is more, small growing follicles with up to three granulosa cell layers are present in women who are relatively gonadotropin deficient with Kallman’s syndrome (5). For practical reasons, information about early follicle development in human ovaries is scarce, although the use of a mouse model carrying mutations for both SCID and hpg has shown that follicles in human xenografts do not develop beyond the two-layer stage (6). This implies that follicle growth initiation is independent of FSH, although this assumption requires verification.

The FSH receptor belongs to a superfamily of receptors characterized by their interaction with the intracellular G proteins. This family of receptors share a similar topology, namely they span the cell membrane 7 times, with an extracellular N-terminus and an intracellular C-terminus (7). The FSH receptor gene (FSHr) has recently been sequenced in several species, including the human (8, 9, 10), and has been found to consist of 10 exons, with exons 1–9 coding the extracellular N-terminal domain and exon 10 coding the transmembrane and C-terminal domains (11).

This progress now provides an opportunity to study the ontogeny of FSHr using a nested PCR protocol of sufficient sensitivity and specificity for analyzing single follicles isolated from the human ovary. Although a positive result would not prove functional significance, it would at least lay the basis of a better understanding of the physiology of early follicle development.

	Materials and Methods

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Tissue collection

Ovarian biopsies measuring approximately 1 mm thick were collected from 11 women undergoing cesarean section for elective indications. This work was approved by the research ethics committee of the Leeds General Infirmary. Tissue was chopped into fragments 2 x 2 mm and transferred to tubes containing Leibovitz L-15 medium plus 1 mg/mL collagenase type IA and 8 U/mL of deoxyribonuclease I (Sigma Chemical Co, St. Louis, MO) and incubated at 37 C for 2 h to partially disaggregate, as described previously (12). Follicles were dissected free of stroma, and the stage of follicle development was confirmed under x250–400 magnification, using an inverted microscope with Hoffman optics (Olympus, Tokyo, Japan). Stages of development were characterized as illustrated in Fig. 1, and those with more than two layers of granulosa cells were defined as multilaminar follicles.

View larger version (80K): [in this window] [in a new window]   Figure 1. Isolated human ovarian follicles. A, Primordial follicle, a small oocyte surrounded by single layer of pregranulosa cells; B, primary follicle with one cell enlarged (early primary); C, primary follicle with a single layer of enlarged granulosa cells; D, two-layer follicle; E, three-layer multilaminar (preantral) follicle; F, large multilaminar (preantral) follicle. Magnification, x300.

RNA extraction and reverse transcription

Cells were suspended in phosphate-buffered saline and centrifuged in a microfuge for 10 min at 4 C; then the process was repeated. They were resuspended in 100 µL lysis/binding buffer for 2 min, followed by centrifugation for 1 min at 4 C. The supernatant was added to 150 µg washed Dynabeads (Dynal, Lake Success, NY), and polyadenylated RNA was isolated according to the manufacturer’s instructions (13). Eight microliters of the resulting supernatant fluid were used for reverse transcription, and the remainder was stored at -70 C to serve as a negative control for PCR. In some primordial follicles, smaller messenger RNA (mRNA) elution volumes were used to allow all the mRNA to be reverse transcribed and tested for low level FSHr expression. In these instances, mRNA controls could not be used.

Reverse transcription was performed using a first strand cDNA synthesis kit (Pharmacia, St. Albans, UK), and the cDNA was stored at -70 C to be used within 12 h for the first round of PCR.

Nested PCR

Oligonucleotide primers were designed on the basis of published FSHr sequences (16) and commercially synthesized (Leeds University Biotechnology, Leeds, UK). The first round of PCR was performed with a sense (5')-primer derived from nucleotides 632–651 in exon 8 (5'-ATGATGTTTTCCACGGAGCC-3') and an antisense primer (3') representing nucleotides 1092–1111 in exon 10 (5'-ACCATATCAGGACTCTGAGG-3'). These primers spanned introns 8 and 9, which precludes amplification of genomic DNA based on the intron sizes in the rat (15 and 3 kilobases, respectively) (12). Nesting was performed by combining a sense primer representing nucleotides 776–795 (5'-AAAAGCTTGTCGCCCTCATG-3') with the antisense primer used in primary reaction spanning intron 9. Both rounds of PCR consisted of an initial 5-min denaturation at 95 C followed by 30 cycles of 95 C for 1 min, 50 C for 1 min, and 72 C for 2 min. A final extension step of 72 C for 5 min was included for both PCRs. To confirm the presence of cDNA in the PCR reaction, a human ß-actin control was run in parallel with each sample. The 20-µL PCR reaction mixture contained 2 µL cDNA, buffer, 200 µmol/L deoxy-NTPs, 1.5 mmol/L Mg²⁺, 0.5 U Taq polymerase (Promega Corp., Southampton, UK), and 0.5 pmol/µL primers. For nesting, 1 mL of a 1:50 dilution of primary PCR product was used. As negative controls, nonreverse transcribed mRNA, rectus abdominus muscle cDNA, stromal tissue cDNA, as well as DNA-free samples were run. Granulosa cells aspirated during egg collection for in vitro fertilization were used as positive controls. To check the sensitivity of the protocol, all cDNA prepared from some follicles was used in the first round of PCR. This represents a 15-fold increase in the amount of mRNA used in the PCR in these cases. PCR products were viewed after separation on 1% agarose gel. To confirm the identity of the bands, the PCR products were cloned and sequenced using automated fluorescent methods (ABI PRISM Automated Dye Terminator Cycle Sequencing, Perkin-Elmer, Warrington, UK).

	Results

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Tissues were collected from 11 patients, ranging from 25–33 yr in age. Between 1–5 follicles were tested from each patient (median = 3) from a total of 35 follicles available for analysis. The results were similar in the patients, with 9 having at least 1 growing follicle expressing the FSHr gene. Three follicles that were negative for FSHr were excluded from the study because ß-actin mRNA could not be detected. A nested PCR on mature granulosa cells aspirated during in vitro fertilization procedures yielded a 479-bp (first round) and a 335-bp product (nesting), confirming the functionality of the primers. Rectus muscle (Fig. 2, lane 6), stromal tissue (Fig. 2, lane 5), nonreverse transcribed follicular mRNA, and DNA-free controls were uniformly negative, and granulosa cell controls from antral follicles were always positive (Fig. 2, lane 1) as anticipated. Microscopic examination confirmed the follicle stage, that each follicle was free of adhering tissue or other follicles, and that transfer to the RNA extraction tube was successful.

View larger version (57K): [in this window] [in a new window]   Figure 2. FSH receptor mRNA expression in isolated human primary follicles. Lane 1, Mature granulosa cells (positive control); lane 2, primordial follicle; lanes 3 and 4, primary follicles; lane 5, ovarian stroma; lane 6, rectus abdominus muscle. m, Marker; a, actin primers; f, FSHr primers. Both primary follicles and the granulosa cells from mature antral follicles were positive for FSHr. Primordial follicle, ovarian stromal cell, and rectus muscle samples were negative for FSHr and positive for ß-actin.

The results are summarized in Table 1. They show that although none of the 9 primordial follicles expressed the FSHr, 33% of the primary and 2-layer follicle stages were positive. Among the 12 primary follicles, 2 had only 1 enlarged granulosa cell (early primary follicle stage), and these were both negative for FSHr mRNA. Positive results at this stage were obtained from follicles that at least contained multiple cuboidal granulosa cells, albeit not invariably. Even at the 2-layer stage, only one third of the follicles were positive for FSHr mRNA. In contrast to the variable expression at earlier growing stages, FSHr mRNA was detected in all multilaminar preantral follicles. The overall frequency of FSHr expression differed significantly between groups (by ² test, P = 0.01), and the higher incidence of positive results among the growing follicles than at the primordial follicle stage was statistically significant (by Fisher’s exact test, P < 0.05).

	Discussion

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The follicles were collected during late pregnancy. Follicle growth initiation and preantral follicle growth continue uninterrupted from late intrauterine life through menopause (1). Thus, there is no evidence that either FSH responsiveness or FSHr gene expression is altered during pregnancy.

This is the first study to indicate that the FSHr gene is not expressed in primordial follicles until after they enter their growth phase. FSHr expression may be triggered at a specific morphological stage in granulosa cells, such as when they change shape or, as present results suggest, may occur progressively during the transition from primordial to primary and two-layer stage. Interestingly, FSHr parallels expression of the gene for the sperm receptor, zp3 (17), and they are two of the earliest signs that follicles have left their dormant state.

The absence of FSHr gene at the primordial stage is an indication that FSH probably does not play a role in the process of growth initiation, and the search for the responsible factor(s) must turn elsewhere. It does not, however, rule out the possibility of an indirect effect mediated by factors from larger follicles. Expression of FSHr in the primary follicle (although variably at that stage) could imply that these follicles are responsive to the hormone, though this requires verification by testing for the protein and the ability to respond to FSH by stimulating cAMP production. Unfortunately, even short term maintenance of primordial and primary follicles has proved challenging as these follicles disintegrate rapidly in vitro (our unpublished observations). The FSHr protein expression could also not be studied because Western blot analysis is too insensitive to study individual follicles. It will be interesting to carry out an immunocytochemical analysis of the FSHr, although negative results would not necessarily indicate an absence of translation, as this technique is less sensitive than PCR.

There is, however, in vitro evidence with human preantral follicles that FSH has a stimulatory effect on multilaminar-preantral follicles. In a study of human preantral follicles cultured for up to 96 h, FSH increased the uptake of [³H]thymidine, implying that in vitro growth had been stimulated (18). In another study, mice carrying mutations at both the SCID and hpg loci (and, therefore, both immunodeficient and hypogonadotropic) were given xenografts of human ovarian tissue. Follicles evidently initiated growth, but developed no further than the two-layer stage unless the hosts were injected with purified FSH (6). Likewise, in an in vitro study, bovine primordial follicles began growing in a defined medium in the complete absence of FSH (19). Taken together, these results confirm assumptions based on laboratory rodent studies that follicle growth initiation is gonadotropin independent (20, 21, 22), but shortly afterward, follicular growth is responsive to FSH.

Data in primate species are sparse. Binding of radioiodinated FSH was detected in multilaminar and antral follicles of adult monkey ovaries and in fetal ovaries during advanced stages of gestation (23, 24), but negative results were obtained during the first and second trimesters in human fetal ovaries (24), which indicates that prefollicular germ cells do not express FSHr. Contrary to much of the evidence, a recent study of human ovaries and fallopian tubes identified weakly positive signals for the FSHr mRNA by in situ hybridization in pregranulosa cells of primordial follicles as well as a stronger signal in preantral growing stages (25). No signal was found in oocytes. Unfortunately, strict comparisons between the studies are invalid because the primordial follicle stage was not well defined, nor were the numbers of observations reported or the hybridization results illustrated. Another study claimed on the basis of in situ hybridization evidence of 1000 primordial follicles in sheep ovaries that the FSHr was definitely absent (26), and yet another study suggested that FSHr and insulin-like growth factor I and insulin-like growth factor II genes are coexpressed in small preantral follicles in the porcine ovary (27). Even so, we should interpret evidence from such techniques and hormone binding studies with caution, because it is difficult to locate the origin of signals emanating from radioactive decay with sufficient precision to be sure whether it is in the diminutive pregranulosa cells. Nested PCR analysis of isolated follicles is a more sensitive technique and does not suffer from this disadvantage.

The absence of FSHr gene expression in primordial follicles proves that elevated serum FSH levels, as a result of either ovulation induction or perimenopausal changes (28), cannot directly accelerate follicle depletion. On the other hand, its presence in single layer primary follicles indicates that FSH may play a much earlier role in human follicular development than had been suspected and raises the possibility that the stimulatory effects of ovulation induction agents are not limited to antral follicles. The biology of the early follicular stages deserves more attention than it has received hitherto, and any evidence of heterogeneity in FSHr and response to hormones at these early stages is of particular interest, as it may set the stage for differential follicular growth, which is central to the emergence of the dominant follicle (29).

	Footnotes

 
¹ This work was supported by a Royal College of Obstetricians and Gynecologists-American College of Obstetricians and Gynecologists Fellowship (to K.O.) and a research grant from the Special Trustees of the United Leeds Teaching Hospitals National Health Service Trust (to R.G.G.).

Received February 27, 1997.

Accepted July 15, 1997.

	References

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