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1 Department of Farm Animal Health, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands
2 Faculty of Veterinary Medicine, PPGCV, State University of Ceara, Fortaleza, CE, Brazil
(Requests for offprints should be addressed to J R V Silva, Department of Farm Animal Health, Faculty of Veterinary Medicine, Utrecht University, PO Box 80151, 3584 CL, Yalelaan 7, Utrecht, The Netherlands; Email: j.r.v.silva{at}vet.uu.nl)
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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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Activin was originally identified as a peptide growth factor from the ovarian follicular fluid that stimulates FSH secretion in cultured anterior pituitary cells (Vale et al. 1986). It is a homodimer or heterodimer of two similar but distinct ß subunits (ßA and ßB). The dimerisation of activin ß subunits gives rise to three forms of activin, that is, activin A (ßAßA), activin B (ßBßB) and activin AB (ßAßB). Both ßA and ßB subunits can also combine with the 
subunit and form inhibin-A (ßA) and -B (ßB) respectively. In the ovary, activin-A is the most studied type (Tisdall et al. 1994, Yokota et al. 1997, Zhao et al. 2001, Thomas et al. 2003) and plays an important role in ovarian follicle development (Ohshima et al. 2002, Lovell et al. 2003), but activin-B has also been described (Bristol & Woodruff 2004). The activities of activin are modulated by its binding protein follistatin, which was originally purified from mammalian follicular fluid as a FSH inhibitor (Phillips & de Kretser 1998). Follistatin binds activin with high affinity, and its binding effectively neutralises the bioactivities of activin in a variety of target tissues (Knight & Glister 2001, Fisher et al. 2003).
Like most of the TGFß superfamily members, activin signals through two types of closely related receptors designated type I and type II, each represented by two isoforms, that is, activin receptor types IA (ActR-IA), IB (ActR-IB), IIA (ActR-IIA) and IIB (ActR-IIB). Activin first binds to a type II receptor, which in turn recruits and activates a type I receptor by phosphorylation. The activated complex of activin and its receptors then stimulates the downstream intracellular signalling molecules that are translocated to the nucleus to regulate target gene transcription (Pangas & Woodruff 2000). ActR-IB is the predominant type I receptor for activin, while ActR-IA might have specificity for either activin or bone morpho-genetic protein signals (Massagué & Chen 2000). In vitro studies have demonstrated that activin-A stimulates pre-antral follicle development in bovine (Hulshof et al. 1997) and rodent isolated follicles (Liu et al. 1998, Smitz et al. 1998, Zhao et al. 2001), increases FSH receptor and FSH-induced luteinising hormone (LH) receptor production in rat granulosa cells (Minegishi et al. 1999, Tsuchiya et al. 1999) and suppresses androgen production in human theca cells (Hillier et al. 1991). Activin also regulates ovarian steroidogenesis in primates (Alak et al. 1998) and oocyte maturation and developmental competence in cattle (Silva & Knight 1998). In contrast, there are also reports that activin-A has no effect on bovine primary follicle development (Fortune 2003) and oocyte maturation in the rat (Tsafiri et al. 1989) and pig (Coskun & Lin 1994).
Expression of protein and mRNA for activin-A and activin receptors in the ovary has been localised in both oocyte and granulosa cells of follicles at various developmental stages (rodents: Wu et al. 1994, Zhao et al. 2001; primates: Roberts et al. 1993, Sidis et al. 1998; pigs: van den Hurk & van de Pavert 2001; and ruminants: Tisdall et al. 1994, Hulshof et al. 1997, Izadyar et al. 1998). In addition, activin-A and activin receptors have been detected in human theca cells (Pangas et al. 2002) and porcine corpora lutea (van den Hurk & van de Pavert 2001). Interestingly, follistatin has also been localised in granulosa cells of rat (Nakatani et al. 1991), sheep (Tisdall et al. 1994) and man (Roberts et al. 1993), and in oocytes, granulosa cells and corpora lutea of cattle (Izadyar et al. 1998, Singh & Adams 1998). Although there is convincing evidence that activin signalling is important for ovarian function, information on its localisation and function is mainly obtained from rodents and cows. Furthermore, most of these studies emphasise only antral follicles, and some of them show conflicting results. To improve our knowledge about the factors that control ovarian follicle development in mammals and to explore possible physiological differences, it is very important to conduct studies with other species, such as goats. Goats are present on all continents and are commercially seen as highly attractive livestock, since they constitute an important source of products such as meat, milk, fibre and skin.
The present study was carried out to examine the possible existence of an intrafollicular activin–follistatin–activin receptor system and its distribution in ovaries of cyclic goats to find evidence of a possible role of this system in ovarian activity, folliculogenesis especially. Therefore, mRNA expression of activin-A (ßA subunit), its type I and type II receptors and its binding protein follistatin was investigated by reverse-transcription polymerase chain reaction (RT-PCR), while the presence and cellular localisation of the corresponding proteins were analysed by immunohistochemistry.
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Materials and Methods |
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During the breeding season, ovaries (n=56) were recovered from slaughtered cyclic adult mixed-breed goats and transported to the laboratory in a thermos flask within 1 h. Sixteen of those ovaries were fixed overnight at room temperature in 4% (w/v) paraformaldehyde in PBS (pH 7.4), and subsequently dehydrated and embedded in paraffin wax (Histoplast; Shandon Scientific Ltd, Pittsburgh, PA, USA) in preparation for immunohistochemical studies. The remaining 40 ovaries were used to recover cells and tissues for RT-PCR.
Immunohistochemistry
Localisation of activin-A, follistatin and ActR-IIA/B and -IA was performed on serial 5-µm sections cut from ovaries of eight different goats. These sections were mounted on poly-L-lysine-coated slides, dried overnight at 37 °C, deparaffinised in xylene and rehydrated in a graded ethanol series. Endogenous peroxidase was blocked by incubating the deparaffinised sections in 3% hydrogen peroxide in methanol for 10 min. The sections were then washed with PBS, and the epitopes were activated by microwaving the sections for 7 min at 900 W in 0.01 M citrate buffer (pH 6.0). After microwave treatment, the sections were washed in PBS/0.05% Tween (PBS-T; Merck, Darmstadt, Germany) before being incubated for 30 min with 5% normal goat serum in PBS to minimise non-specific binding. The primary antibodies were as follows: 1) rabbit antiactivin-A (Innogenetics, Ghent, Belgium); 2) mouse antifollistatin (R&D System Europe, Abingdon, UK); 3) rabbit anti-Act-RIIA that cross-reacts with ActR-IIB (Celgen, Leuven, Belgium) and mouse anti-ActR-IA (R&D System Europe). All antibodies were diluted 1:50. Unfortunately, the available anti-ActR-IB is not suitable for use in goat tissue. The sections were incubated overnight at 4 °C in appropriate dilutions of the antibodies. All other incubations and washes were performed at room temperature. After incubation with an antibody, sections were washed three times with PBS-T and incubated for 45 min with biotinylated secondary antibody (goat antirabbit or goat antimouse IgG from Vector Laboratories, Burlingame, CA, USA), diluted 1:200 in PBS containing 5% normal goat serum. Next, the sections were washed three times in PBS-T before being incubated for 45 min with an avidin–biotin complex (1:600) (Vectastain Elite ABC kits; Vector Laboratories). The sections were then washed three times in PBS and stained with diaminobenzidine (DAB) (0.05% DAB in Tris/HCl, pH 7.6, and 0.03% H2O2 – Sigma tablets) until a precipitate formed or for a maximum of 20 min. The stained sections were rinsed in PBS and water, and counterstained for 10 s in Mayer’s haematoxylin. Finally, the sections were washed for 10 min in running tap water, and subsequently dehydrated in a graded ethanol series followed by xylene treatment and mounting in Pertex (Cellpath Ltd, Hemel Hempstead, UK). The staining intensity was scored as follows: absent (–), weak (+), moderate (++), or strong (+++). Sections were analysed by two independent researchers. Controls for non-specific staining were performed by (1) replacing the primary antibody with IgGs from the same species in which the specific antibody was raised, at the same concentration, and (2) incubation with diaminobenzidine reagent alone to exclude the possibility of non-suppressed endogenous peroxidase activity.
Classification and measurement of follicles
Ovarian follicles were classified as 1) primordial (one layer of flattened/cuboidal granulosa cells), 2) primary (a single layer of cuboidal granulosa cells), 3) secondary (two or more layers of cuboidal granulosa cells), 4) small antral follicles (<3 mm in diameter, with multiple granulosa cells enclosing an antrum), and 5) large antral follicles (3–6 mm). The diameter of follicles was calculated by the method described by Van den Hurk et al.(1994).
Collection of cells and tissues for RT-PCR
The ovaries were rinsed in saline (0.9% NaCl) containing antibiotics (100 IU/ml penicillin and 100 µg/ml streptomycin), after which 10 of them were used for isolation of primordial, primary and secondary follicles. The remaining ovaries were used for collection of oocytes, cumulus cells, mural granulosa cells and thecal cells from small and large antral follicles; and samples of corpora lutea and ovarian surface epithelium.
Early-stage follicles, that is, primordial, primary and secondary, were isolated by the mechanical procedure described previously (Lucci et al. 1999). After isolation, these follicles were washed several times to remove the stromal cells completely, and then placed by category into separate Eppendorf tubes in groups of 15. This procedure was completed within 2 h, and all samples were stored at –80 °C until the RNA was extracted. In previous work from our group, histological analysis was performed to confirm goat preantral follicle classification after isolation (Lucci et al. 1999).
From a second group of ovaries (n=20), cumulus–oocyte complexes (COCs) were aspirated from small (1–3 mm) and large (3–6 mm) antral follicles with an 18-gauge needle attached to a tube in line with a vacuum pump. From the follicle content so collected, compact COCs were selected as described by Van Tol & Bevers (1998). Thereafter, the cumulus was separated from the oocyte by a combination of vortexing and aspiration via a narrow-bore Pasteur pipette. Denuded oocytes, and cumulus and mural granulosa cells were separated, washed four times in PBS, packed in tubes in groups of either 10 denuded oocytes, cumulus cells from 10 COCs or samples of mural granulosa, and stored at –80 °C until RNA extraction.
To collect theca cells, small (n=10) and large antral follicles (n=10) were isolated from ovaries (n=5) and dissected free of stromal tissue with forceps, as described previously (Van Tol & Bevers 1998). The follicles were then bisected, and the granulosa cells were scraped off with a scalpel. Next, the theca cell layers were vortexed for 1 min in 1 ml HEPES-buffered M199 (Gibco BRL, Paisley, UK) supplemented with penicillin/streptomycin, transferred to a fresh 1 ml of buffer, vortexed again for 1 min, washed twice in 2 ml HEPES-buffered M199, collected and stored at –80 °C. From another group of ovaries (n=5), small pieces of corpus luteum and surface epithelium were collected and stored at –80 °C until RNA extraction. Three samples of each tissue sample were analysed.
Extraction of total RNA and reverse transcription
Isolation of total RNA combined with on-column DNAse digestion was performed with the RNeasy mini kit and the RNAse-free DNAse set (Qiagen). According to the manufacturer’s instructions, 350 µl lysis buffer were added to each frozen sample, and the lysate was aspirated through a 20-gauge needle before being centrifuged at 10 000 g for 3 min at room temperature. The lysates of theca cells, corpus luteum and ovarian surface samples were then subjected to proteinase K treatment (6.7 mAU/ml, Qiagen) at 55 °C for 10 min. Thereafter, all lysates were diluted 1:1 with 70% ethanol and introduced to a mini-column. After binding of the RNA to the column, DNA digestion was performed with RNAse-free DNAse (340 Kunitz units/ml) for 15 min at room temperature. After washing of the column three times, the RNA was eluted with 30 µl RNAse-free water.
Prior to the reverse-transcription reaction, the eluted RNA samples were incubated for 5 min at 70 °C, and chilled on ice. Reverse transcription was then performed in a total volume of 20 µl made up of 10 µl sample RNA, 4 µl 5X reverse transcriptase buffer (Gibco BRL, Breda, The Netherlands), 8 units RNAsin, 150 units Superscript II reverse transcriptase (Gibco BRL), 0.036 U random primers (Life Technologies BV, Leiden, The Netherlands) and 10 mM DTT and 0.5 mM of each dNTP. The mixture was incubated for 1 h at 42 °C and for 5 min at 80 °C, and then stored at –20 °C. Minus RT blanks were prepared under the same conditions, but without inclusion of reverse transcriptase.
Amplification of cDNA
PCR reactions were carried out in 200 µl tubes (Biozym, Landgraaf, The Netherlands), using 1 µl cDNA as template in 25 µl of a mixture of 2 mM MgCl2, 200 µM of each dNTP and 0.5 µM each of primers, and 0.625 units Taq DNA polymerase (HotStarTaq; Qiagen). The primers used for amplification of the inhibin/activin ßA subunit, follistatin, Act-RIIA, -RIIB, -RIA, and -RIB, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH), are presented in Table 1
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A standard sequencing procedure (ABI PRISM 310 Genetic analyzer, Applied Biosystems) was used to verify the specificity of the PCR products.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Activin-A and follistatin proteins were detected in oocytes of primordial follicle stage onward (Fig. 1A and J), and in granulosa cells of primary (Fig. 1B and K) and secondary follicles (Fig. 1C and L). Occasionally, activin-A was found in granulosa cells of primordial follicles (Fig. 1A), but no reaction to either activin-A or follistatin was found in theca cells from secondary follicles (Fig. 1C and L). ActR-IIA/B and ActR-IA proteins were found in oocyte and granulosa cells of primordial (Fig. 2A and J), primary (Fig. 2B and K) and secondary follicles (Fig. 2C and L), but not in theca cells of these follicles.
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) and follistatin (Fig. 1M–O), except for small antral follicles that had a weak reaction to activin-A (Fig. 1D, Table 2). Additionally, these same sites had a positive reaction to both ActR-IIA/B (Fig. 2D–F) and ActR-IA (Fig. 2M–O), except the theca cells of large antral follicles, which did not react to ActR-IA (Fig. 2O). Apart from follicles, immunoreactions to activin-A, follistatin, ActR-IIA/B and ActR-IA were observed in corpora lutea (Fig. 1G and P) and ovarian surface epithelium (Fig. 1H and Q). The relative intensity of immunohistochemical staining for activin-A, follistatin, ActR-IIA/B and ActR-IA in all cell types studied is illustrated in Table 2. For all antibodies tested, control reactions (Figs 1I and R and 2I and R) confirmed the absence of non-specific staining.
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Amplification of cDNA from primordial, primary and secondary follicles resulted in specific products for inhibin/activin ßA subunit, follistatin, ActR-IIA, -IA and -IB (Fig. 3). Transcripts for both the ßA subunit of activin (653 bp) and follistatin (566 bp), and all activin receptors (IIA (527 bp), IIB (184 bp), IA (431 bp) and IB (315 bp)) were also detected in cDNA from oocytes, cumulus, mural granulosa and theca cells collected from small and large antral follicles as well as from corpus luteum and ovarian surface epithelium (Fig. 3). The expression of the housekeeping gene (GAPDH) is also illustrated in Fig. 3. Sequence analysis of the amplified ßA subunit of activin, follistatin, and ActR-IIA, -IIB, -IA and -IB products confirmed their specificity. Amplification of –RT (without reverse transcriptase) or water blanks (cDNA replaced by water) controls yielded no specific products in any of the reactions.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In caprine antral follicles, both activin-A and follistatin proteins were present in the oocyte, cumulus cells, mural granulosa cells and theca cells. These compounds apparently are formed in these cells, since their mRNA was also detected at the same locations. Transcription of ßA subunit can also form inhibin-A, but this will occur only when there is an excess of ßA subunit production (Findlay 1993, Knight & Glister 2001). The role of inhibin on folliculogenesis has been recently reviewed by Knight & Glister 2001. Activin-A protein has been detected in oocytes and granulosa cells in rat (Ogawa et al. 1991), human (Yamoto et al. 1992, Roberts et al. 1993, Wada et al. 1996), bovine (Izadyar et al. 1998, Silva et al. 2003) and porcine (van den Hurk & van de Pavert 2001) antral follicles. In man (Roberts et al. 1993) and pig (van den Hurk & van de Pavert 2001), theca cells of antral follicles also immuno-reacted to the activin-A protein. Thus far, expression of activin-A mRNA in antral follicles has appeared to be restricted to the granulosa cells in rat (Meunier et al. 1988), primate (Roberts et al. 1993), sheep (Braw-Tal 1994, Tisdall et al. 1994) and pig (van den Hurk & van de Pavert 2001), while in cows it was also detected in oocytes (Izadyar et al. 1998). Follistatin protein and mRNA have been demonstrated in oocyte and granulosa cells from bovine antral follicles (Izadyar et al. 1998, Silva et al. 2003). Expression of follistatin mRNA in antral follicles has also been reported in rat (Nakatani et al. 1991), man (Roberts et al. 1993, Sidis et al. 1998) and sheep (Braw-Tal 1994, Tisdall et al. 1994), and it appeared to be confined to granulosa cells.
We furthermore demonstrated ActR-IIA/B and ActR-IA proteins and the mRNAs for ActR-IIA, -IIB, -IA and -IB in oocytes, cumulus cells, mural granulosa cells and theca cells in the goat. Previously, our group reported the expression of ActR-IIA mRNA and protein in oocytes and granulosa cells of pig (van den Hurk & van de Pavert 2001) and cow (Izadyar et al. 1998) antral follicles, while in those of the human, ActR-IIA and -IIB were detected in granulosa cells and theca cells (Pangas et al. 2002). In mouse and human antral follicles, mRNAs for all four activin receptor subtypes were expressed in both oocytes and granulosa cells (Sidis et al. 1998). In vitro, activin-A stimulated antrum formation (rat: Zhao et al. 2001), FSH receptor and FSH-induced LH receptor production (rat: Minegishi et al. 1999, Tsuchiya et al. 1999), cytochrome P-450 aromatase activity, and thus oestrogen synthesis, in granulosa cells (human: Mukasa et al. 2003). Activin also suppressed androgen production in theca cells from different species (human: Hillier et al. 1991; bovine: Wrathall & Knight 1995; and ovine: Campbell & Baird 2001). Consequently, activin is thought to play a crucial role in basal growth, recruitment and selection of antral follicles through stimulation of proliferation and FSH receptor expression in granulosa cells and modulation of steroidogenesis in granulosa and theca cells, its actions being time and concentration dependent and regulated by follistatin (reviewed by Findlay et al. 1993, 2002, Driancourt 2001, Knight & Glister 2001). Activin-A is also involved in the regulation of oocyte maturation in several species (bovine: Silva & Knight 1998; human: Alak et al. 1998; and mouse: Sidis et al. 1998). The distribution pattern of activin-A, its binding protein follistatin and activin receptors in goat antral follicles points to an important role of these proteins in antral follicle development in this species.
Apart from follicles, we found evidence for the presence of an activin–follistatin–activin receptor system in corpora lutea and ovarian surface epithelium, because of the presence of both mRNA and protein for activin-A, follistatin and all four types of activin receptors at these sites. With regard to corpora lutea, expression of protein and mRNA for activin-A and follistatin has previously been demonstrated in man (Roberts et al. 1993, Wada et al. 1996) and cow (Singh & Adams 1998). Both Act-RIIA protein and mRNA were detected in pig luteal tissue (van den Hurk & van de Pavert 2001), and, in vitro, activin-A suppressed progesterone production by primate luteal cells (Brannian et al. 1992). Expression of activin-A protein and mRNA in ovarian surface epithelium has previously been demonstrated in pig (van den Hurk & van de Pavert 2001) and cow (Hulshof et al. 1997). Choi et al.(2001) demonstrated that activin inhibits growth and induces apoptosis in cultured ovarian surface epithelial cells. As mentioned above, the activin–follistatin system has been demonstrated to control growth and differentiation of antral follicles by affecting gonadotrophin receptor synthesis and steroidogenesis. The activin–follistatin system presence in goat corpora lutea and surface epithelium may therefore point to a regulatory function in or fine tuning of their activity.
In conclusion, activin-A, its binding protein follistatin and its receptors are formed in all types of goat follicles and in antral follicles in all compartments. In addition, they are generated in corpora lutea and ovarian surface epithelium. The widespread distribution pattern of the follistatin–activin–activin receptor system in goat ovaries points to a crucial role of this system in various reproductive processes, including follicle growth and differentiation, and luteal activity.
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Acknowledgements |
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References |
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Received 24 March 2004
Accepted 21 July 2004
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