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REPRODUCTION-DEVELOPMENT |
Prince Henry’s Institute of Medical Research, Clayton, Victoria 3168, Australia
Address all correspondence and requests for reprints to: Ann Drummond, Ph.D., Prince Henry’s Institute of Medical Research, Clayton, Victoria 3168, Australia. E-mail: . ann.drummond{at}med.monash.edu.au
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Members of the TGFß family of peptides, including activin, transduce their signals via serine/threonine kinases, of which there are 2 types (5, 6); I and II, the type I receptors being the primary transducers of signals. Activin receptors comprise type IA (ActRIA) and IB (ActRIB), IIA (ActRIIA) and IIB (ActRIIB), of which there are four forms designated ActRIIB1–4 (7, 8). Activin receptors have been identified in rat, human, and mouse ovaries (9, 10, 11, 12, 13) at various developmental stages, although their presence and pattern of expression in the rat ovary during the establishment of folliculogenesis has yet to be determined. Functional antagonism of the activin signaling pathway has recently been demonstrated by the inhibin coreceptor ßglycan (14). In the presence of ßglycan, the affinity of inhibin for the activin type II receptor is enhanced leading to sequestration of the receptor and an inability of activin to bind and initiate signal transduction. ßglycan may therefore be important in determining whether activin signaling components interact to form functional pathways that may have special significance in ovarian cells. Expression of ßglycan mRNA by the ovary has not been previously reported, although protein has been localized to the rat ovary in a preliminary study (14).
Following ligand binding, the type II receptor kinases phosphorylate and thereby activate type I receptors, which subsequently phosphorylate and activate the appropriate intracellular substrates or Smad(s) (reviewed in Ref. 15). Smads 1, 5, and 8 are phosphorylated by activated bone morphogenetic protein (BMP) receptors (16, 17, 18), and Smads 2 and 3 are phosphorylated by activin and TGFß receptors (19, 20). A complex with Smad 4 is then formed and these composites translocate to the nucleus to activate the transcription of target genes (for review, see Ref. 15). Inhibitory Smads (Smad 6 and 7) have been identified (21, 22), although no definitive mechanism of action has been elucidated. They have been shown to associate with type I receptors, effectively blocking the association and activation of Smads (23), inhibit phosphorylation of receptor-Smad complexes (22) and compete with Smad 4 for binding to the receptor activated complex (24). Whatever the mechanism of Smad 6 and Smad 7 action, the endpoint is a block of signal transduction. Little information is available on the expression of the intracellular signaling Smad proteins by the ovary. Smad 2 protein has been localized to hen granulosa cells (25) and human oocytes and Smad 2 mRNA detected in human oocytes (26). Smad 3 mRNA has been detected in mouse granulosa cells (27).
It is conceivable that members of the TGFß superfamily other than activin may play a role in the acquisition of gonadotropin sensitivity in the ovary. Recently, BMP-4 and -7 have been shown to increase the sensitivity of isolated granulosa cells to FSH stimulation (28), as has TGFß in the past (29, 30). Given our desire to elucidate the roles of activin and other members of the TGFß superfamily in establishing and perpetuating the folliculogenic process, we investigated the expression of the activin receptor subtypes, ßglycan and Smads, in the ovary of the postnatal rat. This model has proven valuable in our studies of folliculogenesis (1, 31, 32) because defined follicle populations appear for the first time, in a sequential manner, during the first 12 d after birth. Thus, the specific aims of the study were to: 1) establish the presence of mRNA for the activin/BMP receptor subtypes, ßglycan and Smads in the postnatal ovary; 2) determine the pattern of expression of these mRNAs during this developmental period and establish if there is coordinated expression of receptors with specific Smads; and 3) localize activin receptor and Smad proteins to specific ovarian cells.
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Materials and Methods |
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RNA extraction
Ovaries were dissected free of fat and adhering tissue and homogenized in 1 ml Ultraspec RNA reagent (Biotecx: Fisher Biotec, Melbourne, Australia). After 5 min on ice, 0.2 ml chloroform per ml of Ultraspec RNA reagent was added to the samples, which were then shaken vigorously and stored at 4 C for 5 min before centrifuging for 15 min at 12,000 x g. RNA was precipitated from the aqueous phase with 1 volume of isopropanol, after which the pellet was washed twice with ethanol, air-dried, and resuspended in sterile water. To ensure that the RNA was completely dissolved, the samples were incubated for 10 min at 60 C. At least three independent pools of RNA were prepared from the ovaries of postnatal rats at each age studied. The number of ovaries/pool ranged from 24–40, depending on the age of the animal.
RT
RNA (2 µg) was reverse transcribed with 50 U Moloney murine leukemia virus reverse transcriptase (Expand, Roche, Sydney, Australia) and final concentrations of 1x cDNA synthesis buffer (supplied with enzyme), 1 mM deoxynucleotide triphosphates (Roche), 20 U Rnasin (Promega Corp., Sydney, Australia) 10 mM dithiothreitol and 25 pmol oligo deoxythymidine-15 (Roche), as previously described (31).
Real time PCR
mRNA expression was analyzed using the Roche LightCycler (Roche, Mannheim, Germany) as previously described (32). In these studies, two separate rat ovarian cDNAs prepared from the ovaries of 22-d-old rats, were assigned arbitrary unitage and employed as the assay standard and quality control, respectively. For PCR, 2 µl each of the standard cDNA pool diluted 1:2, 1:20, 1:200, and 1:2000 (arbitrarily designated 1.0, 0.1, 0.01, and 0.001, respectively), the quality control cDNA pool diluted 1:25, and the sample cDNAs diluted 1:10–1:20 in sterile water, were added to individual capillaries. Taq enzyme, deoxynucleotide triphosphates, reaction buffer, and SYBR GREEN I dye were supplied in the FastStart DNA Master SYBR Green I kit (Roche, Mannheim, Germany), of which 2 µl/capillary was added. Primer concentrations of 17.5–25 pmol were added to each capillary. The primer specific nucleotide locations, sequences and product sizes are shown in Table 1
. Magnesium concentrations (2–4 mM), annealing temperatures (57-64 C), and extension times (number of seconds = product size/25 plus 3) were determined for individual primer sets. The capillary volume was made up to 20 µl with sterile water. Forty cycles of PCR were programmed to ensure the threshold crossing point (cycle number) was attained. Fluorescence emission was monitored continuously during cycling. At the completion of cycling, melting curve analysis was carried out to establish the specificity of the amplicons produced. In addition, each amplicon was sequenced to verify the identity of the amplified product (data not shown). The level of expression of each mRNA and their estimated crossing points in each sample were determined relative to the standard preparation using the LightCycler computer software. A ratio of specific mRNA/GAPDH amplification was then calculated, to correct for any differences in efficiency at RT. The relative abundance of the mRNAs, expressed as fold changes, was extrapolated from crossing point data. A difference of 1 PCR cycle in crossing point number translates into a 2-fold change in mRNA expression. In many cases, an estimated 2-fold change in mRNA abundance was not statistically different from the appropriate control, given the variation in crossing point number between individual pools.
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Immunohistochemistry
Antibodies directed against ActRIA, ActRIB, ActRIIA, ActRIIB, ßglycan, (R&D Systems, Minneapolis, MN) Smad 2, 4, 5, and 8 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), were used to localize the respective proteins to 5-µm sections of formalin-fixed paraffin-embedded ovary using standard immunohistochemical protocols. These antisera were selected because they were suitable for localizing rat proteins and had previously been used for Western blotting and/or immunohistochemistry (14, 33, 34, 35). Briefly, sections were dewaxed in histosol (Australian Biostain, Australia), dehydrated in ethanol, and washed in water. The sections were microwaved in 0.1 M citrate buffer, pH 7.4, on high power for 12 min and allowed to cool before equilibrating in 0.1 M PBS, pH 7.4. The sections were blocked for 30 min in 1% blocking reagent (Roche), after which the primary antibody, appropriately diluted 1:10–1:200 (0.02–50 µg/ml) was added and the sections incubated for 48 h at 4 C. Following extensive washing in PBS, the biotinylated second antibody (DAKO Corp., Carpinteria, CA) was added to the sections for a 60-min incubation at room temperature. After washing in PBS, the sections were incubated with a peroxidase conjugated avidin-biotin complex (Vector Elite, Vector Laboratories, Inc., Burlingame, CA) for 60 min at room temperature after which the reaction product was developed using 3,3' diaminobenzidine tetrahydrochloride (DAKO Corp.) and hydrogen peroxide in PBS. The sections were counterstained with hematoxylin, dehydrated in ethanol, cleared in histosol and coverslips mounted using DPX (British Drug House, Poole, UK). Control sections received either buffer or IgG (goat) diluted appropriately, in place of primary antibody. The ovaries of 6–10 rats, two sections/ovary were examined for each antibody in three separate experiments. The staining was scored by two independent observers, as outlined in the legends to the tables. Follicles were classified as follows: primordial follicles contained an oocyte surrounded by a single layer of flattened squamous granulosa cells; primary follicles contained an oocyte surrounded by a single layer of cuboidal granulosa cells; secondary follicles contained an oocyte surrounded by two or more layers of granulosa cells, and antral follicles were those in which the oocyte was contained within multiple layers of granulosa cells in which fluid filled spaces were apparent.
Statistical analysis
Data collected from at least three separate RT-PCR experiments were analyzed using GBstat. Statistical significance was determined by ANOVA in conjunction with a posthoc multiple comparison test (Fisher’s least significant difference test). P values of < 0.05 compared with the appropriate control were regarded as statistically significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). The expression of all four activin receptor mRNAs was highest at d 4. Thereafter, the expression of ActRIB and ActRIIA mRNAs declined and remained at this level up to d 12. ActRIA and ActRIIB mRNA expression were also reduced at d 8, but by d 12 had returned to the level of expression recorded in ovaries of 4-d-old rats. ßglycan mRNA was expressed by postnatal rat ovaries from 4-, 8- and 12-d-old rats (Fig. 1). The level of expression was similar in ovaries of 4- and 8-d-old rats, but by d 12 ßglycan mRNA expression had increased 2-fold relative to d 4 levels.
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and 3A). In ovaries of 4- and 8-d-old rats, similar levels of expression were noted for ActRIA, ActRIB, and ActRIIA, whereas ActRIIB was the least abundant receptor mRNA (Tables 2 and 3A). By d 12, the expression of ActRIB mRNA had declined to levels observed for ActRIIB (Table 3A).
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). The ovarian expression of Smad 3, 6, and 8 mRNAs did not change significantly between d 4 and 12. The mRNA expression of the remaining Smads (Smads 1, 2, 4, 5, 6, and 7) was highest at d 4, reduced in comparison at d 8 and by d 12, either remained at the level of expression recorded at d 8 (Smad 4 and 7), continued to decline (Smad 5), returned to d 4 levels (Smad 2), or assumed a level of expression in between d 4 and d 8 (Smad 1) (Fig. 2).
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and 3B). Consistently, the difference in expression of these Smads was separated by 6–7 PCR cycles (Table 2), which translates into 64- to 128-fold more Smad 2 than Smad 3, given that 1 PCR cycle represents a 2-fold change in mRNA (Table 3B). The common Smad 4 and Smad 2 showed a similar pattern of expression except that Smad 2 was more abundant than Smad 4 (Tables 2 and 3B). Smad 8 was the least abundant BMP-related Smad in the ovary during d 4–12, whereas Smad 5 was the most abundant mRNA in 4- and 8-d-old ovaries. By d 12, Smad 5 was expressed at levels similar to Smad 1 (Tables 2 and 3C). Smad 4 mRNA was expressed at levels similar to Smad 1 during d 4–12 (Tables 2 and 3C). The inhibitory Smads 6 and 7 were expressed at similar levels in the ovary during d 4–12 (Table 2), although this level of expression was 4- to 8-fold less than Smad 4 mRNA abundance (Tables 2 and 3D).
Immunohistochemical localization of ActRIA, ActRIB, ActRIIA, ActRIIB, ßglycan, and Smads 2, 4, 5, and 8 to postnatal rat ovaries
Protein for all four activin receptors, Smads 2, 4, 5, and 8 and ßglycan were immunolocalized to the ovaries of 4-, 8-, and 12-d-old rats (Figs. 3 and 4, and Tables 4–6). Activin receptor proteins were essentially confined to oocytes and granulosa cells, except ActRIB, which was also present in theca cells of secondary and antral follicles. In sections of d 4 ovaries, ActRIA, ActRIB, ActRIIA, and ActRIIB were localized to the oocyte cytoplasm of primordial follicles with granulosa cells expressing only ActRIB and ActRIIA and then only in a few follicles (Table 4). We did not detect protein for any of the activin receptors in primary follicles on d 4. In sections of d 8 ovaries, primordial (oocytes), primary (granulosa cells and oocytes) and secondary follicles (oocytes and granulosa cells) localized most of the activin receptor proteins (Fig. 3, A–D, and Table 5). Expression of ActRIA protein by granulosa cells was not observed until follicles reached the secondary stage and ActRIIB proteins were detected in oocytes of primordial and primary, but not secondary follicles. A similar pattern of expression was observed in sections of d 12 ovary (Table 6). Activin receptor and ßglycan proteins were not localized to blood vessels. ßglycan protein was localized to primordial, primary, secondary, and antral follicle oocytes and granulosa cells and theca cells during the postnatal period (Fig. 3E, and Tables 4–6). Cytoplasmic and nuclear staining patterns were evident in most cases. ßglycan protein was not detected in blood vessels.
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and Tables 4–6) and primary follicles (oocytes and granulosa cells) of 4-d-old rat ovaries, with both nuclear and cytoplasmic staining patterns (Table 4). Sections of 8- and 12-d-old ovaries localized Smad 2 protein to primordial (granulosa cells), primary (oocytes and granulosa cells), secondary and antral follicles (oocytes and granulosa cells), and theca (Fig. 4A, and Tables 5 and 6). Smad 4 protein was localized to primordial follicle oocytes (but not granulosa cells) and primary follicles (oocytes and granulosa cells) of 4-d-old rat ovaries (Table 4). In addition to these follicle populations, the ovaries of 8- and 12-d-old rats also localized protein to secondary follicles and theca cells (Fig. 4B, and Tables 5 and 6) with a predominantly cytoplasmic staining pattern. Smad 5 and Smad 8 proteins were localized to the oocyte cytoplasm of primordial follicles (d 4–12), primary follicles (d 4–12), and secondary and antral follicles (d 8 and 12) (Fig. 4, C–D, and Tables 4–6), with an unusual half-moon staining pattern evident in most oocytes, particularly small follicles (Fig. 4, C–D). From the intensity of staining, Smad 5 appeared to be more abundant than Smad 8. Smad 5 protein was also present in granulosa cells of primary follicles of d 4 ovaries but was not detected in granulosa cells of any other follicles at any other time (Fig. 4 C–D). Smad 8 protein was not detected in granulosa cells or theca cells at any stage. |
Discussion |
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Activin/BMP receptor mRNA expression
The expression of activin receptor mRNAs is consistent with previously published reports indicating that human cumulus cells, granulosa-luteal cells and oocytes and mouse oocytes and cumulus-oocyte complexes contain ActRIA, ActRIB, ActRIIA, and ActRIIB (10, 12, 13, 36). Activin subunit mRNA has also been localized to human and mouse oocytes (12). In the rat ovary, He et al. (37), reported that ActRIA mRNA was mainly localized to oocytes of preantral and antral follicles, whereas ActRIB mRNA was not detected using the in situ hybridization technique employed. More recently, Zhao et al. (38) reported that oocytes and somatic cells of rat preantral follicles expressed ActRIIA and localized activin A protein. Our data suggests that ActRIB mRNA is present in the postnatal rat ovary (presence of protein confirmed immunohistochemically), but the level of expression declined with increasing follicular development. ActRIIA and ActRIIB mRNAs are expressed by the ovaries of d 15 rat embryos (39) and the ovaries of immature and adult rats (9, 11, 40). The presence of ActRIIB mRNA in the ovary has been somewhat equivocal given that Cameron et al. (11) reported low level expression in oocytes and granulosa cells by in situ hybridization, whereas Aloi et al. (40) did not detect any ActRIIB mRNA in adult ovary by RT-PCR. Our studies indicate that postnatal rat ovaries (and immature 25-d-old rat ovaries, unpublished data) express ActRIIB mRNA and protein, but it was clearly the least abundant of the activin receptor subtypes. It is important to keep in mind that BMPs can also use receptors initially identified as activin receptors, for signaling (18, 41).
Regulation of activin receptor mRNAs
Little is known about what regulates the expression of activin receptor mRNAs in the ovary. In adult cycling rats, ActRIA and ActRIIA have been shown to be regulated by gonadotropins and E2 (40). ActRIA and ActRIIA expression declined in vivo after the proestrous gonadotropin surge, or in response to the administration of human CG and FSH to hypophysectomized rats. E2 decreased ActRIA and increased ActRIIA mRNA in ovaries of hypophysectomized rats. In the presence of both gonadotropins and E2, ActRIA expression was reduced, the authors concluding that this response was mediated by E2 produced in response to FSH stimulation. ActRIIA expression was inhibited by gonadotropins and E2, essentially mimicking the effect of gonadotropins alone, indicating a direct action of gonadotropins potent enough to counteract the stimulatory effect of E2. Immediately after birth, the rat ovary is unresponsive to FSH stimulation. By postnatal d 8, secondary (preantral) follicles are present and capable of responding to FSH, thus the decline in ActRIIA mRNA expression we report here, may be mediated by FSH. The production of E2 by follicles increases with development so that antral follicles, present in ovaries of 12-d-old rats, have the largest capacity. We cannot say at this time whether E2 influences the expression of ActRIA and ActRIIA by the postnatal rat ovary. Similarly, the regulation of ovarian ActRIB, ActRIIB, ßglycan, and Smad mRNAs has yet to be addressed.
Protein localization
Our data indicate that a functional activin signaling pathway exists in the oocytes of primordial follicles from d 8 onwards, which is consistent with a role for activin in oocyte maturation that has previously been reported for human (42), bovine (43), rat (44, 45), zebrafish (46, 47), and primate (48) oocytes. While activin receptor and Smad 4 proteins were present in primordial follicle oocytes of ovaries from 4-d-old rats, Smad 2 protein was not detected and therefore the ability of activin to transduce a signal in these oocytes is questionable. It is possible that the expression of Smad 2 was simply below the level of sensitivity of the immunohistochemistry protocol. Alternatively, Smad 3 may be involved in the transmission of activin signals, although this seems doubtful given that Smad 3 mRNA was clearly the least abundant of the activin/TGFß Smads. In contrast, granulosa cells of primordial follicles across the postnatal period failed to consistently express any of the proteins under investigation, suggesting that granulosa cells of these early stage follicles could not support an action of activin/TGFß.
An interesting finding of these studies was the apparent age-dependent appearance of activin receptors by primary follicles. Oocytes and granulosa cells of primary follicles from 4-d-old rat ovaries did not localize activin receptor proteins. These same follicle populations in ovaries of 8- and 12-d-old rats, however, did contain detectable activin receptor proteins. We noted that most of the primary follicles present at d 4 contained granulosa cells, which were enlarged and transparent, leading us to hypothesize that these follicles exist in a transitional state and may not express differentiated parameters, e.g. activin receptors, during this transformation. Our earlier studies (1) clearly demonstrated an effect of activin on inhibin production by d 4 ovarian cells in culture, the implication being that activin receptors are present and that granulosa cells of primary follicles were the likely source of inhibin. If this is indeed the case, then perhaps activin receptor proteins are expressed at a level below the sensitivity of our immunohistochemistry protocol. Alternatively, after 2 d in culture, our ovarian cells may have matured, acquired activin receptors and responded to activin stimulation (in this instance by producing inhibin). This would be consistent with time course studies undertaken in our laboratory, which indicate that ovarian cells from 4-d-old rats produce most of their inhibin 24–48 h (32 ; Drummond, A. E., M. Dyson, and J. K. Findlay, unpublished observations) after initiating the culture.
ßglycan in the ovary
ßglycan mRNA expression was stable in the ovaries of 4- and 8-d-old rats, in which oocytes and granulosa cells of early stage follicles predominate. Between d 8 and d 12, a significant increase in expression occurred, which coincided with the appearance of theca cells around secondary preantral follicles. Consistent with these results, ßglycan protein was localized to oocytes, granulosa cells, and theca cells as expected. Given the duality of ßglycan’s roles, its ability to act as an inhibin coreceptor and to functionally antagonize activin signaling (14), the localization of ßglycan to specific cellular populations will be paramount in ascertaining its role. Inhibin has been shown to stimulate human thecal cell steroidogenesis (49), and in this capacity ßglycan may act as an inhibin coreceptor. The picture is more complex in regards to granulosa cells as these produce inhibin and activin, respond to activin and express activin receptors, thus inhibin through ßglycan, may act to antagonize activin action in these cells.
BMP signaling
An action of BMP-4 and -7 on steroidogenesis has recently been described (28), with the data prompting the authors to propose that BMPs may act as luteinization inhibitors. In some respects, notably the enhancement of E2 production by FSH-stimulated granulosa cells in culture, the action of these BMPs is similar to activin (28) raising issues of redundancy and relative importance of individual factors in ovarian regulation. The similarity of signal transduction pathways and confusion in regards to receptor activation by these ligands has not clarified matters. The complexity of the receptor-signaling pathway is highlighted by reports that BMP-7 and activin bind to the same type II receptors (18, 41) but recruit distinct type I receptors. Activin has been shown to recruit ActRIB and transduce signals via Smad 2 or 3, whereas BMP-7 recruits ActRIA and transduces signals through Smad 1, 5, and 8 (18). Our data show that ActRIA protein is confined to oocytes at all stages and granulosa cells of secondary and antral follicles, suggesting that these are primary sites of BMP action. Localization of Smad 5 and 8 proteins to oocytes (of most follicles) is also consistent with an action of BMPs. Given that Smad 8, was clearly the least abundant of the BMP-related Smads and its mRNA expression did not change with follicular status, we hypothesize that Smad 1or 5 mediate BMP signaling in the oocyte. We have yet to localize Smad 1 protein to ovarian cells. It is unclear as to whether the location of the Smad proteins within the cell is an indicator of activation, i.e. does nuclear localization of protein indicate an activated Smad? In these studies, both cytoplasmic and nuclear localization were evident for Smad 2 and 4, although not always simultaneously within the same cells or in the same follicle populations. The localization of Smad 8 to the cytoplasm of oocytes and Smad 5 to oocyte and granulosa cell cytoplasm would, if this premise is correct, suggest inactive proteins and a nonfunctional signaling pathway.
In conclusion, we have established the presence of activin receptor, Smad and ßglycan mRNAs in postnatal rat ovaries and localized proteins indicative of activin/TGFß and BMP signaling pathways to specific ovarian cells (Table 7). The role of activin in promoting follicular differentiation (1, 50) is strengthened by the colocalization of important signaling elements to growing follicles and those in which an antrum is forming. A direct action of activin on oocytes is also supported by the data. It is likely that an important role for inhibin in the ovary is to antagonize activin action, via ßglycan, so that follicles capable of responding to activin and producing inhibin and ßglycan may be destined for atresia. The presence of BMP signaling elements in oocytes suggests an involvement of BMPs in oocyte maturation, in addition to their postulated role as luteinization inhibitors (28). Additional studies are required to establish whether individual follicle populations localize unique signaling pathways or if active pathways of the TGFß family can be identified in follicles. The spatial (cell specific) and temporal expression patterns of the components of these pathways will determine the capacity of follicular cells to respond and the type of response induced by members of the TGFß family, if and when they are expressed.
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Acknowledgments |
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Footnotes |
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Abbreviations: ActRIA, ActRIB, ActRIIA, and ActRIIB, Activin/BMP receptors; BMP, bone morphogenetic protein.
Received October 4, 2001.
Accepted for publication December 14, 2001.
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References |
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-subunit levels in granulosa and thecal layers of developing preovulatory follicles in the chicken. Endocrinology 139:1164–1171 and ß mRNA expression: impact of development and estrogen. Mol Cell Endocrinol 149:153–161[CrossRef][Medline]
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) point to granulosa cells that are in the process of differentiating. Arrows (
) indicate half-moon staining pattern (oocyte cytoplasm) observed with antibodies for Smad 5 (C) and 8 (D). x20 Magnification.