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a Animal Reproduction and Biotechnology Laboratory, Department of Physiology, Colorado State University, Fort Collins, Colorado 80523
b AgResearch, Wallaceville Animal Research Centre, Upper Hutt, New Zealand
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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1 over noncarriers (++) [1]. In contrast, animals homozygous for the FecXI gene (II) have "streak" gonads and are sterile [2]. Fetal II sheep demonstrate normal germ cell development and follicular formation until approximately Day 100 of gestation. However, as ovarian follicles start to grow (i.e., from Day 100 onward), development beyond the primary stage is impaired, and normal secondary follicles are not observed [3, 4]. Disruption of normal folliculogenesis at this stage of development appears to be due to a lack of granulosa cell proliferation as oocytes increase in diameter to >40 µm [3, 5]. As a result, ovaries of II sheep contain many abnormal follicles characterized by an enlarged oocyte and lacking a normal complement of granulosa cells. In addition, follicles with oocytes at various stages of degeneration and oocyte-free follicles, or "nodules," are observed [3, 4]. Thus, compared to other genotypes, it seems that a fundamental event in early follicular growth is impaired in II animals during late fetal, neonatal, and adult life. Whether the FecXI mutation affects oocyte development, granulosa cell differentiation, or both is not known [3]. Thus, studies designed to examine the cellular and intraovarian effects of this gene have the potential to provide insight into the mechanisms controlling initiation of folliculogenesis and oocyte development. The signal(s) and mechanism(s) that regulate gonadotropin-independent folliculogenesis are unknown. Although the potential role of a number of growth factors and their receptors in preantral follicular growth has been examined, the majority of these factors are not expressed until after follicular growth has been initiated (for review see [6, 7]). Recently, a novel member of the transforming growth factor ß superfamily, termed growth and differentiation factor-9 (GDF-9), was shown to be expressed in oocytes of domestic ruminants from the primordial follicle stage onward [8]. The observed oocyte specificity and timing of GDF-9 expression is consistent with the concept that this growth factor may be important in initiation of folliculogenesis in these species; however, the precise role of GDF-9 in gonadotropin-independent folliculogenesis in sheep remains obscure.
Interestingly, GDF-9-deficient mice have a phenotype similar to that in II sheep. For example, follicular development is arrested at the primary follicle stage, associated granulosa cells fail to proliferate, and antral cavities and thecal cell layers are not formed [9, 10]. Furthermore, lack of GDF-9 leads to an abnormal increase in oocyte size, with eventual loss of the oocyte and formation of steroidogenic granulosa cell clusters [11]. Collectively, these data suggest that GDF-9 and the FecXI gene product play fundamental roles in the regulation of preantral follicular growth and oocyte development. Although GDF-9 has been mapped to chromosome 5 in sheep [12] and, therefore, is not a candidate for the Inverdale mutation, it is possible that the FecXI gene could affect GDF-9 expression. Because of the phenotypic similarities between II sheep and GDF-9-deficient mice, we hypothesized that expression of GDF-9 would be inhibited in oocytes in ovaries of II fetal sheep. To test this hypothesis, expression of GDF-9 mRNA in ovaries of II, I+, and ++ sheep at Day 135 of gestation was examined using in situ hybridization.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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All experiments were performed with approval granted from the Animal Ethics Committee of the Invermay Agricultural Centre and the Wallaceville Animal Research Centre, and in accordance with the 1987 Animal Protection (Codes of Ethical Conduct) Regulations of New Zealand.
I+ and ++ female fetuses were obtained by mating progeny-tested I rams (n = 3) with I+ ewes that were themselves derived from progeny-tested I rams. Each fetus was assigned to either the II or I+ genotype from analyses of DNA extracts of muscle tissue recovered from the hind leg. Although the Inverdale genetic mutation is unknown, genotype was determined using three DNA markers (Inv 12, Inv 17, and Inv 07) [13]. Authentic female I+ fetuses were generated by mating progeny-tested rams (n = 2) with ++ ewes. The ++ female fetuses were generated by mating noncarrier rams (n = 3) with ++ ewes.
Fetuses (n = 4 per genotype: II, I+, and ++) were recovered via cesarean section at 135 days of gestation (day of mating = Day 0). Ovaries were removed from fetuses and immersion-fixed in 4% (w:v) phosphate-buffered paraformaldehyde, pH 7.4, for 24 h. Tissue was trimmed and embedded in paraffin (Paraplast; Oxford Laboratory Ware, St. Louis, MO), and serial sections were cut (5–7 µm thick) and transferred to microscope slides coated with 3-aminopropyltriethoxysilane (Aldrich Chemical Company, Milwaukee, WI). Slides were then shipped to the Animal Reproduction and Biotechnology Laboratory at Colorado State University (Fort Collins, CO) for in situ hybridization and examination of mRNA expression.
Use of c-kit and Follistatin
Expression of c-kit mRNA has been localized to oocytes from the primordial (type 1) follicle stage onward in fetal sheep [14], and expression of follistatin mRNA has been detected in granulosa cells of follicles from the secondary (type 3) follicle stage onward in adult sheep [6, 15]. In addition, follistatin mRNA expression has been detected in the oocyte-free follicles, or nodules, of II adult sheep (Dr. J. Juengel, personal communication). Thus, in the present study, in situ hybridizations for c-kit and follistatin served as positive controls for oocyte and granulosa cell expression respectively. Further, follistatin expression in the nodules of II sheep facilitated identification of these structures and, in some cases, allowed for analysis of GDF-9 expression in adjacent sections.
Probe Preparation
A 277-base pair (bp) genomic fragment of ovine GDF-9 exon 2 was cloned into pBS/KS (Stratagene, La Jolla, CA) and digested with either BamHI (T7; antisense) or XbaI (T3; sense) restriction endonucleases for the production of riboprobes [8]. Plasmid DNA (pGEM-3Z; Promega, Madison, WI) containing a 422-bp ovine c-kit cDNA was linearized for the production of riboprobes with either KpnI (SP6; antisense) or PstI (T7; sense) restriction endonucleases [14]. Plasmid DNA (pGEM-3Z; Promega) containing a 726-bp ovine follistatin cDNA was digested with EcoRI (SP6; antisense) or XbaI (T7; sense) for riboprobe production [16].
In Situ Hybridization
Expression of mRNAs encoding ovine GDF-9, c-kit, and follistatin was examined using in situ hybridization as previously described [8, 17] with modifications from Tisdall et al. [15]. Tissue sections (5–7 µm thick) were deparaffinized by immersion in Hemo-De (Fisher Scientific, Pittsburgh, PA) two times for 10 min each time and hydrated by passage through graded alcohols (100–70%) and diethyl pyrocarbonate (DEPC)-treated water for 3 min. Slides were then incubated in 0.2 M HCl for 20 min at room temperature and washed in double-strength SSC (single-strength SSC is 0.15 M sodium chloride, 0.015 M sodium citrate) for 30 min at room temperature. Proteinase K digestion was performed for 5 min at 37°C (proteinase K solution = 2 µg/ml proteinase K [Sigma, St. Louis, MO]; 0.2 M Tris HCl [pH 7.2]; 50 mM EDTA [pH 8.0]). Slides were then dipped in triethanolamine (0.1 M, pH 8.0), incubated in 0.25% acetic anhydride in 0.1 M triethanolamine for 10 min, dipped in double-strength SSC, dehydrated through graded alcohols (50–100%) and air-dried. After linearization of plasmid, antisense and sense 35S[UTP]-labeled riboprobes were generated with the appropriate promoters. Sixty microliters (2 x 107 cpm/ml) of RNA probe in hybridization buffer (for 40 ml: 25 ml deionized formamide, 3.75 ml 4 M NaCl, 0.5 ml 1 M Tris pH 8.0, 1 ml 50-strength Denhardt's solution, 10 ml 50% dextran sulfate, 0.75 ml double-distilled H2O, and 200 mM dithiothreitol) was added to each slide and covered with a glass coverslip. Sections hybridized to GDF-9 and c-kit were incubated overnight in a humidified chamber at 50°C and sections hybridized to follistatin were incubated at 55°C. After hybridization, coverslips were removed by soaking in double-strength SSC. Sections were then washed in double-strength SSC for 10 min, incubated in a ribonuclease (RNase)-A solution (Sigma; 600 µl 10 mg/ml RNase-A into 300 ml double-strength SSC) at 37°C for 30 min, washed in double-strength SSC for 10 min and in a stringent wash solution (0.1-strength SSC, 0.09% ß-mercaptoethanol, 1 mM EDTA) at 55 or 60°C for 2 h (GDF-9 at 55°C; c-kit and follistatin at 60°C), and rinsed twice in 0.5-strength SSC for 10 min. Sections were dehydrated through graded ethanol-ammonium acetate solutions (50–100% ethanols; 0.3 M NH4OAc) and air-dried. Slides were then coated with NTB2 emulsion (Eastman Kodak, Rochester, NY) and stored at 4°C for 22 or 32 days (GDF-9 and c-kit = 32 days; follistatin = 22 days). Emulsion was developed, and tissue sections were stained with hematoxylin and eosin and examined using both brightfield and darkfield optics.
Classification of Follicles
Classification of follicles was based on the scheme developed by Lundy et al. [18]. In this classification system, five well-defined stages of follicular growth from primordial to early antral are recognized. Criteria used to classify ovine follicles into specific developmental stages included the number and appearance of granulosa cells, layers of granulosa cells, and follicle/oocyte diameters. Primordial follicles are classified as type 1 follicles and are characterized by a single layer of squamous granulosa cells. Type 1a follicles are also characterized by a single layer of granulosa cells, but one or more of the granulosa cells surrounding the oocyte are cuboidal. Type 2 follicles are commonly referred to as primary follicles and have 1 to <2 complete layers of cuboidal granulosa cells. Characteristics of subsequent stages using this classification scheme are as follows: type 3, small preantral follicles with 2 to <4 layers of cuboidal granulosa cells; type 4, large preantral follicles with 4–6 layers of cuboidal granulosa cells; and type 5, small antral follicles with >5 layers of granulosa cells and an antral cavity. In the present study, an additional follicle type was added to account for large antral follicles and included all follicles >450 µm in diameter. The percentage of follicles per type that expressed GDF-9, c-kit, or follistatin was determined in all follicles in which the plane of section included the oocyte nucleus. Only nonatretic follicles were considered in the study.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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On Day 135 of gestation, normal follicles from all stages of follicular development (types 1–5) were observed in the ovaries of I+ and ++ sheep. In contrast, normal follicles beyond the type 2 stage were not detected in II sheep. There was no evidence of abnormal follicular structures or abnormal primary follicles in the ovaries of I+ and ++ animals. However, abnormal type 2 follicles characterized by an enlarged oocyte and an abnormal arrangement of granulosa cells were present in all four of the II sheep examined. Furthermore, of the four II animals studied, all had clearly identifiable oocyte-free follicles, or nodules.
Expression of mRNA Encoding GDF-9, c-kit, and Follistatin
In situ hybridization was used to detect follicles that expressed GDF-9, c-kit, and follistatin mRNAs in ovaries of II, I+, and ++ sheep at Day 135 of gestation. The numbers of follicles examined for each follicular class (types 1–5), genotype, and growth factor are summarized in Table 1. In each of the three genotypes, type 1a follicles were the most numerous and ranged from 64–77% of the total number of follicles evaluated.
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Distribution of silver grains over sections of fetal ovaries incubated with the 35S-radiolabeled antisense probe for GDF-9 appeared to be oocyte-specific in all three genotypes (Fig. 1), and oocytes in all follicles from the type 1 follicle stage onward were labeled. Specific hybridization was not evident in any other cell type, including granulosa and theca cells. Enlarged oocytes of abnormal type 2 follicles (total = 21 follicles; Fig. 1A) in the ovaries of II sheep also expressed GDF-9 mRNA, but specific binding of GDF-9 antisense probe to oocyte-free follicles, or nodules, in the ovaries of II fetal sheep was not observed.
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Similar to the case with GDF-9, specific hybridization of the 35S-radiolabeled antisense probe for c-kit was detected in all oocytes from the type 1 follicle stage onward in II, I+, and ++ fetal ovaries (Fig. 2). In addition, c-kit mRNA expression was noted in granulosa cells of type 4 and large antral follicles from I+ and ++ sheep (Fig. 2C). Although granulosa cells in several additional large preantral and antral follicles in the ovaries of I+ and ++ sheep were positive for c-kit expression, the plane of section did not include the nucleus of the oocyte. Therefore, these follicles were not included in data summarized in Table 1. Specific hybridization to other ovarian cell types, including thecal and stromal tissue, was not observed. Although oocytes of abnormal type 2 follicles in the ovaries of II sheep expressed c-kit mRNA (total = 36 follicles; Fig. 2B), specific hybridization to abnormal ovarian structures, including oocyte-free nodules, was not noted (Fig. 2B).
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Expression of follistatin mRNA was restricted to granulosa cells of type 4 and larger follicles in I+ and ++ sheep (Fig. 3B) and was not apparent in oocytes or theca cells at any stage of development examined. Although follistatin expression was detected in granulosa cells of several putative type 3 follicles, the plane of section did not include the oocyte nucleus, and therefore these follicles were not included in the final count. Furthermore, and consistent with the blockage in folliculogenesis at the type 2 stage in II sheep, granulosa cells in follicles from these sheep did not express mRNA for follistatin. Nevertheless, follistatin mRNA expression was detected in the oocyte-free nodules of II sheep (Fig. 3A).
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Specific labeling was not observed in sections incubated with 35S-radiolabeled sense probe for GDF-9 (Fig. 1, B, D, and F), c-kit (Fig. 2D), or follistatin (Fig. 3C).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In domestic ruminants, onset of GDF-9 gene expression begins at the type 1 follicle stage ([8]; present study). This is in contrast to data from mice, in which GDF-9 mRNA expression was not observed until the primary follicle stage [19, 20]. Apparent differences in onset of GDF-9 expression in these species could be due to differences in follicular classification criteria, to sensitivity of the in situ hybridization system used, or to species differences. However, since GDF-9 mRNA expression from the type 1 (primordial) follicle stage has recently been documented in mice [21], it seems likely that discrepancies in GDF-9 mRNA expression between rodents and ruminants are due to procedural differences in the in situ hybridization methodology (e.g., tissue fixation or length of exposure).
In all studies to date, ovarian expression of GDF-9 mRNA has been localized exclusively to oocytes [8, 19–21]. Consistent with this, we did not detect GDF-9 mRNA in oocyte-free follicles, or nodules, in ovaries of II sheep. In contrast, GDF-9 mRNA expression was observed in enlarged oocytes of abnormal type 2 follicles. Thus, despite the putative role of GDF-9 in granulosa cell proliferation and development [9, 11], it would not appear that a lack of GDF-9 gene expression underlies the development of abnormal ovarian structures in II sheep. Clearly, however, we cannot preclude the possibility of a lesion in either GDF-9 protein expression or signaling that may contribute to the abnormal follicular development observed in II sheep.
Specific hybridization for mRNA encoding the tyrosine kinase receptor, c-kit, was localized to oocytes of all follicles from the type 1 stage onward, regardless of genotype. In addition, c-kit mRNA expression was noted in granulosa cells of large preantral and antral follicles in I+ and ++ sheep. Patterns of c-kit mRNA expression in the present study correspond to those reported previously for sheep [14] and are similar to patterns of expression that have been noted in mice [23–25]. Consistent with data reported for adults [6, 15], follistatin mRNA expression was localized to granulosa cells in I+ and ++ sheep but was not detected in granulosa cells of II sheep. Thus, lack of c-kit and follistatin mRNA expression in granulosa cells of II fetal sheep confirms the blockage in folliculogenesis at the type 2 stage of development. In addition, and similar to patterns of expression seen in GDF-9-deficient mice [11], follistatin gene expression was detected in the oocyte-free follicles, or nodules, of II sheep. Expression of follistatin in these structures allowed for positive identification of the nodules and, in some cases, allowed for comparison of GDF-9 mRNA expression in the same structures in nearby sections. Hence, in situ hybridization for c-kit and follistatin mRNAs served as reliable positive controls for both oocyte and granulosa cell gene expression.
The large number of type 1a follicles observed in the present study is consistent with data from cattle [26] and rats [27], in which the majority of follicles had a single layer of granulosa cells with at least one cuboidal cell. Although it is thought that the transition of granulosa cells from squamous to cuboidal is indicative of the initiation of follicular growth, the large number of type 1a follicles in the ovaries of these species suggests that this follicle type does not, in its entirety, represent a transitional stage. Rather, it has been proposed that this follicle type represents a morphologically distinct population of nongrowing primordial follicles together with a portion of follicles developing from the type 1 stage [26–28]. Thus, new markers for the initiation of follicular growth and revision of current follicular classification schemes may be necessary.
In summary, regardless of genotype or stage of development, all oocytes expressed mRNA for GDF-9. On the basis of these results, it appears that phenotypic characteristics of ovaries in II sheep are not due to a lack of expression of mRNA for GDF-9 or alteration in cell- or stage-specific expression of c-kit or follistatin. Although inhibition of GDF-9 gene expression does not appear to be the mechanism of action by which the FecXI gene alters folliculogenesis in II or I+ sheep, the possibility of translation at specific stages of follicular development cannot presently be ruled out. In addition, the FecXI gene may be involved, either directly or indirectly, in regulating expression of receptors for GDF-9. At present, however, neither the FecXI gene product nor the GDF-9 receptor has been isolated or characterized.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by the Colorado State University Agricultural Experiment Station and USDA NRICGP Award Number: 990-2389. K.J.B. was supported by NIH National Research Service Award Number HG07031. 
2 Correspondence: H.R. Sawyer, Animal Reproduction and Biotechnology Laboratory, Foothills Campus, Colorado State University, Fort Collins, CO 80523. FAX: 970 491 3557; hsawyer{at}cvmbs.colostate.edu
Accepted: December 31, 1999.
Received: November 2, 1999.
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