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Endocrinology. Author manuscript; available in PMC 2010 November 3.
Published in final edited form as:
PMCID: PMC2971462

Inhibitor of Differentiation (Id) genes are expressed in the steroidogenic cells of the ovine ovary and are differentially regulated by members of the Transforming Growth Factor (TGF)-β family


Inhibitor of Differentiation (Id) proteins act during embryogenesis and development to repress gene transcription required for lineage commitment, whilst promoting cell growth. Growth factors belonging to the transforming growth factor beta (TGFβ) superfamily of signaling molecules, notably the bone morphogenetic proteins (BMPs) and activin, can regulate Id expression in these tissues. Id expression and function in adult physiology is less well determined and we hypothesized a role for Id proteins in the adult mammalian ovary. Immunohistochemistry for Id1, Id2, Id3 and Id4 in the sheep ovary revealed consistent expression in granulosa and thecal cells of ovarian follicles throughout development. In atretic follicles Id proteins were selectively down-regulated in thecal cells (P<0.0001). Additionally Id1 was universally up-regulated in the cumulus cells adjacent to the oocyte. Immunohistochemistry for phospho (p)-smad 1/5/8 signalling components (stimulated by BMPs) showed a punctate pattern of expression whereas p-smad 2/3 (stimulated by activin) was ubiquitously expressed in follicles. Neither pathway however displayed differential staining in line with Id1 cumulus specific expression, suggesting a more complex relationship between Id1 expression and TGFβ signaling in these cells. Nevertheless, in vitro, stimulation of ovine granulosa cells with BMP6 or activin A led to a respective increase and decrease in Id1 (P<0.0001), Id2 (P<0.0001), Id3 (P<0.0001) and Id4 (P<0.05) transcripts and Id1 gene expression was further manipulated by the oocyte-secreted factors BMP15 and GDF9 (P<0.001). These data confirm that TGFβ signaling can regulate Id gene expression in the sheep ovarian follicle and suggest a functional role for the Id family in the mammalian ovary.

Keywords: Follicle, Granulosa Cells, Bone Morphogenetic Protein, Activin A


The ovary, unlike other adult mammalian tissues, undergoes persistent cyclical remodeling with regulated proliferation, differentiation and cell death. Inhibitors of differentiation (Id) proteins (also known as inhibitors of DNA binding) are a subfamily of regulatory dimeric basic helix loop helix (bHLH) transcription factors. These factors regulate many genes, including those required for growth and differentiation, through binding to E-box (CANNTG) sequences on the promoter region of target genes (1, 2). Id proteins lack a basic DNA-binding domain and can heterodimerize with other bHLH proteins to block chromatin binding and thus subsequent transcriptional activity (3). Four known mammalian Id isoforms (Id1-4) have been identified and can regulate growth and differentiation across embryonic tissues (4-6), where both overlapping and non-redundant functions are reported (7, 8). Predictably, these properties have led to Id proteins being implicated in tumorigenesis in various adult tissues (9, 10).

Upstream mediators of Id expression include members of the transforming growth factor β (TGFβ) superfamily (11), that include the bone morphogenetic proteins (BMPs) and activin (12). Dimeric TGFβ ligands interact with a range of BMP type-1 and type-2 serine/threonine kinase receptor complexes to activate the smad signaling pathway (12). BMPs signal through the smad 1/5/8 pathway whereas activin activates the smad 2/3 cascade (13), and both are antagonised by smad 6 and smad 7 (14). Signaling by TGFβs has been linked to Id gene regulation during development, controling the timing of differentiation and lineage commitment (8, 15-18). This has particular significance for the adult ovary in which BMPs and activin are involved in many processes governing follicle development and oocyte maturation and competency (19-21). Furthermore, although not yet characterized in the adult mammalian ovary, Id mRNA is reported to be expressed in the hen (Gallus domesticus) and the Ids are speculated to be involved in the control and timing of follicle selection and granulosa cell differentiation in this avian species (22, 23).

Although the molecular regulation of proliferation and differentiation in the cycling ovary is not fully understood, members of the TGFβ family appear to have key mechanistic roles. We hypothesized that Id proteins have roles in the regulation of growth and differentiation of the steroidogenic cells of the adult ovary. We therefore studied the localization of Id protein expression (Id1, Id2, Id3 and Id4) in the pre- and post-pubertal sheep ovary. In addition, we hypothesized that Id proteins are differentially regulated by the TGFβ family members through intracellular smad signaling pathways. We therefore localized the smad proteins in the ovine ovary in vivo and investigated the effects of ovarian BMPs (BMP6, BMP15 and/or GDF9) and activin A on the expression of Id genes in primary cultures of non-luteinized granulosa cells in vitro. Herein we report the consistent localization of the Id proteins in the ovine ovary in vivo and their differential gene regulation by BMPs and activin A in vitro.

Materials and Methods


All reagents and chemicals were obtained from Sigma-Aldrich (Poole, Dorset, UK), unless otherwise stated.


Ovaries from Scottish Greyface lambs (n=12), pregnant ewes (n=15) and non-pregnant cycling ewes (n=5), selected from the control cohort of additional studies, were obtained following local ethical committee and regulatory approval. Tissue was fixed in Bouins solution for 24 h, transferred to 70% ethanol and embedded in paraffin wax.

Tissue sections (5μm) were de-waxed and re-hydrated through an alcohol series before undergoing antigen retrieval by pressure cooking in 0.01M sodium citrate buffer pH 6.0 for 5 min. Sections were washed in PBS (2 × 5 min), placed in 3% H2O2 diluted in distilled water for 10 min, followed by PBS washes (2 × 5 min). Tissue was then blocked with 20% normal goat serum (NGS) and 5% bovine serum albumin (BSA) diluted in PBS, for 1 h. Primary antibodies (Table 1) were diluted in blocking solution and incubated with tissue overnight at 4 C. Sections were washed with PBST (PBS +1% tween; 2 × 5 min) before incubation with a biotinylated goat anti-rabbit IgG secondary antibody (Dako, Glostrup, Denmark) at a 1:500 dilution. Following further washes in PBS (2 × 5 min) tissue was incubated with Vectastain ABC Elite tertiary complex (PK-1600 Series, Vector Laboratories, Peterborough, Cambs, UK) for 1 h, and washed in PBS (2 × 5 min) before colorimetric visualization by incubation with 3,3′-diaminobenzidine (DAB; Dako) for 30 sec. Tissue was rinsed in distilled water, de-hydrated in alcohol, counterstained with hematoxylin and mounted.

Table 1
List of primary, secondary and tertiary antibodies used for immunohistochemistry (IH) and immunofluorescence (IF). Secondary antibody; 1) Biotinylated Goat anti-Rabbit IgG (Dako, Glostrup, Denmark) 1:500, 2) Peroxidase Goat anti-Rabbit (Dako) 1:200. Tertiary ...

Primary antibodies were incubated with blocking peptide, prior to application, or in the absence of a specific blocking peptide (smad 6 and smad 7) negative controls consisted of incubation with non-specific rabbit immunoglobulins and omission of the primary antibody. To ensure the binding specificity of smad 6 and smad 7, Western blotting was additionally carried out as previously described (24), using protein extracted from ovine granulosa cells. Specific bands were visualized for smad 6 and smad 7 at approximately 52 and 42 KDa, respectively, as anticipated (data not shown). Histological images were captured using an Olympus Provis BX2 microscope (Olympus America Inc. Center Valley, PA, USA) equipped with a Canon E0S 30D Microcam camera (Canon Inc Headquarters, Tokyo, Japan).


Sheep ovary sections were treated in the same way as for immunohistochemistry but for the following alterations to the protocol. Following incubation with the secondary antibody on day 2 and washing, the appropriate tertiary antibody was applied (Table 1) for 1 h if using streptavidin or 10 min if using tyramide detection systems. Sections were subsequently retrieved by microwaving for 2.5 min in 0.01M sodium citrate buffer pH 6.0 (Tyramide detection system only) and then blocked in NGS for 1 h before overnight incubation with the second primary antibody at 4 C. On day 3 the appropriate secondary and tertiary complexes were applied as described above and the sections were counterstained with 4′,6-diamidino-2-phenylindole (DAPI; Invitrogen, Paisley, UK) diluted 1:1000 in PBS, for 10 min, before mounting with PermaFluor (Thermo Fisher Scientific, Cheshire, UK). Images were captured using a LSM 510 Meta Confocal microscope and Zen 2008 Software (Carl Zeiss Ltd., UK).

Quantification of thecal Id expression in healthy and atretic follicles

To obtain a quantitative measurement of thecal Id expression in follicles, staining intensity for Id1, Id2, Id3 and Id4 proteins were classified in three groups: absent, partial or intense stain. Follicles were classified as atretic or non-atretic based on immunostaining for activated caspase-3 in a serial section. Forty non-atretic and forty atretic follicles (50:50 pre-antral and antral) were examined across 6-8 different ovaries and a count performed categorizing follicles into the appropriate staining class based on clear visual reference pictures agreed by three observers. Proportional data were presented as a percentage of the total number of follicles examined in each group, and examined using a chi-squared test.

Cell culture and quantitative real time (qRT)-PCR

Granulosa cells were collected and cultured as previously described (25). Briefly, follicles (<3.5mm) were collected from ovaries of ten Scottish Greyface ewes during the estrous cycle. Follicles were hemisected and the granulosa cells collected by flushing thecal shells using a 1mL syringe. The supernatant containing granulosa cells was removed and cells collected by centrifugation, washed, and re-suspended in culture medium (McCoys 5a medium with sodium bicarbonate, supplemented with 0.1% BSA, 0.5X PenStrep, 3mM L-glutamine, 5μM transferrin, 0.3mM testosterone, 4nM selenium, 0.01μM insulin, 0.01μM ovine IGF-1 LR3 (Novozymes Biopharna AU Ltd, Australia), and 1nM FSH (NIDDK). Three separate experiments consisted of pooling granulosa cells from 3-4 animals to obtain sufficient numbers. Around 75,000 granulosa cells per well were cultured in 200μL medium with the addition of 100ng/mL activin A and/or BMP6, BMP15 and/or GDF9 (R&D Systems) in treatment wells, or carrier-only in control wells, at 37 C/5% CO2 for 24 h. These concentrations were chosen based on the existing literature regarding BMP and activin effects on ovine steroidogenic cells (26, 27).

Cells were lysed and RNA was extracted using the Qiagen RNeasy Micro Kit (Qiagen Ltd., West Sussex, UK). Lysed cells form 2-3 wells were pooled in order to obtain adequate RNA concentration and purity (A260/A280 ratio) which were measured using a NanoDrop 1000 spectrophotometer (Thermo Fisher Scientific). RNA was stored at −80 C until cDNA was synthesized from 200ng total RNA per reaction using the High Capacity cDNA reverse transcription kit (Applied BioSystems, CA, USA) and thereafter stored at −20 C.

Primer sets were designed for amplification of partial regions of target genes by qRT-PCR (Table 2). Primers were pre-validated in the sheep using conventional PCR and DNA sequencing was performed to confirm the authenticity of the product. Further to this primer efficiency was tested by generating standard curves (cDNA diluted 1:2, 1:4, 1:8 and 1:16) in qRT-PCR reactions. A 10μL final reaction volume was prepared using 2X PowerSYBR Green PCR Master Mix (5μL; Applied BioSystems), 5μM primer pairs (0.5μL), cDNA (1μL) and nuclease free water. qRT-PCR cycling programme consisted of a denaturing step (95°C for 10 min), annealing and extension step (95°C for 15 sec, 60°C for 1 min), repeated 40 times, and a dissociation step (95°C, 60°C and 95°C for 15 sec each). Each reaction was carried out in duplicate. Negative controls included a reaction using cDNA prepared with the omission of reverse transcriptase, and a reaction substituting cDNA with nuclease free water. The relative expression level of each target gene to GAPDH was quantified using the delta delta Ct method and the control data standardized between runs. Data is presented as the mean ± sem and statistical analysis was performed using a one way ANOVA with Bonferroni pair-wise comparison following logarithmic transformation. P values of P<0.05 were regarded as significant.

Table 2
List of primer sequences used for qRT-PCR and amplicon size.


Id proteins are expressed in ovine follicles

Id proteins were localized in sheep ovaries from different functional stages including pre-pubertal, pregnant and non-pregnant states. Specific immunostaining was observed for Id1, Id2, Id3 and Id4 in the follicles of all ovaries examined (Fig. 1). These were consistently localized to granulosa cells in follicles throughout development from primary to pre-ovulatory stages. In addition, consistent weaker staining was observed in the theca cells of these follicles (Fig. 1). Id2 immunostaining was also apparent in the oocytes at all follicular stages and there was strong antibody binding observed across the ovarian stroma (Fig. 1d-f). Weak Id3 expression was visible in the oocytes of primordial follicles (Fig. 1g; black arrows) whereas the zona pellucida exhibited a high level of Id3 expression across follicle development (Fig 1h and i). Specific immunostaining was also observed in endothelial cells for each Id protein examined (Fig. 1), particularly Id3 where there was strong blood vessel localization of this protein (Fig. 1g; grey arrows).

Figure 1
Immunohistochemistry for Id1 (a-c), Id2 (d-f), Id3 (g-i) and Id4 (j-l) protein (brown) in the sheep ovary, illustrates from left to right, staining in primordial or primary, secondary or early antral, and tertiary or late antral follicles. Arrow in b) ...

Differential localization of Ids in granulosa cells

The Id1 antibody revealed intense staining of peri-oocytic cumulus cells (Fig. 1a-c), a striking observation that was reproduced across follicular development and in all specimens, including pre-pubertal lamb and pregnant sheep ovaries as well as cycling ovaries. Although Id4 was more intensely immunostained than Id3 their localization was similar across the granulosa cells and was maintained throughout follicle development (Fig. 1g-l). However, unlike Id1, Id3 and Id4, immunoreactivity for Id2 was consistently most intense in those mural granulosa immediately adjacent the antral cavity of tertiary follicles (Fig. 1f).

Id immunostaining is reduced in the theca cells of atretic follicles

There was differential staining of Id proteins in the theca cells of some follicles (Fig. 2a). Id1, Id2, Id3 and Id4 protein expression was reduced in the thecal layer of atretic follicles, by reference to activated caspase-3 immunostaining in serial sections (Fig. 2c), while granulosa cell expression was maintained (Fig. 2a and b). This phenomenon was encountered consistently in atretic follicles and could be quantified so that intense antibody staining in the thecal layer of healthy follicles was lost resulting in partial or absent expression of Id1, Id2, Id3 and Id4 proteins in follicles undergoing atresia (Fig. 2d-g; P<0.0001). No significant differences for Id protein expression, or alteration in thecal cells in atresia, were observed between pre-antral and antral follicles (Fig. 2).

Figure 2
Id2 expression in adjacent healthy and atretic follicles at low (a) and high (b) magnification and activated caspase-3 protein expression (c) in the corresponding ovine follicles. Arrows in a) show thecal layer and in c) depict positive caspase immunoreactivity ...

Lack of specific peri-oocyte phospho (p)-smad immunostaining in ovine follicles

The most striking feature of Id localization in the ovine ovary was the intense staining for Id1 in the cumulus granulosa cells closest to the oocyte. We therefore hypothesized that an oocyte-derived paracrine factor may regulate the expression of Ids in the neighboring cells. As members of the TGFβ superfamily are excellent candidates for an oocyte-secreted factor we investigated the localization of the main TGFβ signaling pathways in the follicles. P-smad 1/5/8 proteins could be localized to the nucleus of both theca and granulosa cells (Fig. 3a and b). Indeed, the punctate pattern of more intense staining (Fig. 3b) suggests a specific regulated role for this pathway in such cells. However the staining was not increased or decreased in the granulosa cells next to the oocyte in any follicle examined. The alternative p-smad 2/3 pathway could also be detected in nuclei of both theca and granulosa cells of the follicle (Fig. 3d and e). Although this staining was more uniform it also did not demonstrate a differential staining pattern in the granulosa cells surrounding the oocyte. Although it is clear that the cells surrounding the oocyte operate both smad 1/5/8 and 2/3 pathways the consistent immunostaining pattern did not reveal a direct relationship with the peri-oocyte Id1 expression in cumulus granulosa cells.

Figure 3
Immunohistochemistry (brown) for p-smad 1/5/8 (a and b), p-smad 2/3 (d and e), smad 6 (g and h) and smad 7 (j and k) in the sheep ovary. Arrow in g) identifies smad 6 positive immunostaining in granulosa cells and in h) indicates staining in a blood vessel. ...

Peri-oocyte localization of smad 6 in granulosa cells

Both of the inhibitory smads, smad 6 and smad 7, were found to have widespread ovarian expression (Fig. 3). Smad 6 revealed a similar localization to smad 1/5/8 where staining was limited to some granulosa and thecal cells only (Fig. 3g and h). However more intense nuclear staining was observed in some cumulus cells, notably around the oocyte, and also in cortical blood vessels (arrows in Fig. 3g and h, respectively). Smad 7 staining was more uniform in the granulosa and thecal cells of all of the follicles examined (Fig. 3j and k). To ensure the specificity of smad 6 and 7 antibody binding in the sheep, Western blotting for these proteins was also performed in ovine granulosa cell protein extracts. Specific bands were visualized for both smad 6 and 7 at the expected molecular weight confirming the expression of these proteins in this cell population and supporting the specificity of the immunohistochemistry.

The relationship of the smads to Id1 was subsequently investigated using co-immunofluorescence (Fig. 4). P-smad 1/5/8 did not co-localize in the cumulus layer with Id1 (Fig. 4a). There was however some co-localization between smad 6 and Id1 staining in the peri-oocyte granulosa cells (Fig. 4b). This suggests that manipulation of the smad signaling system may yet have a role in the specific staining pattern of Id1 in the peri-oocyte granulosa cells.

Figure 4
Co-immunofluorescence of Id1 (red) with p-smad 1/5/8 (green) (a) and smad 6 (green) (b) in follicles of the sheep ovary. Merged pictures include DAPI nuclear counterstain (blue). Scale bars = 100μm.

Activin and BMP signaling regulates Id gene expression in ovine granulosa cells in vitro

In order to investigate the effect of stimulating the smad 2/3 and smad 1/5/8 pathways on Id expression, ovine granulosa cells were cultured with activin A and/or BMP6 and the levels of Id gene mRNA were analysed by qRT-PCR. Overall ANOVA analysis revealed that Id1 (P<0.0001), Id2 (P<0.0001), Id3 (P<0.0001) and Id4 (P<0.05) were all changed by the treatments in the same pattern (Fig. 5). Stimulation with activin A led to the down-regulation of Id1 and Id3 mRNA expression whereas conversely, BMP6 treatment up-regulated these genes (Fig. 5; Bonferroni pair-wise analysis P<0.001). It was particularly notable for Id1 expression where activin A and BMP6 treatment led to a 5-fold decrease and 7-fold increase in mRNA, respectively (Fig. 5a). Furthermore, treatment with activin A significantly negated the BMP6-induced up-regulation of the Id1 (Fig. 5a; Bonferroni pair-wise analysis P<0.05).

Figure 5
Effect of activin A and/or BMP6 (a), and GDF-9 and/or BMP15 (b) on Id1 gene expression, and activin A and/or BMP6 on Id2 (c), Id3 (d) and Id4 (e) gene expression, in ovine granulosa cells cultured for 24 h. Values are mean ± sem of relative mRNA ...

To further explore the potential roles of TGFβ mediated regulation of Id1 expression in the cumulus cells, an experiment was conducted where the granulosa cell population received doses of oocycte-secreted factors BMP15 and/or GDF9 and Id1 gene expression was measured (Fig. 5.b; ANOVA P<0.001). Similarly to the effects produced with the BMP6 cultures, BMP15 treatment also led to an increase in Id1 transcript (Fig. 5b; Bonferroni pair-wise analysis P<0.01). GDF9 alone did not significantly alter Id1 gene expression however in combination with BMP15 this increased 8-fold (Fig. 5b; Bonferroni pair-wise analysis P<0.001).


This study describes Id protein expression across the developing mammalian ovary and provides evidence for functional roles in normal adult tissue. We have shown for the first time that Id1, Id2, Id3 and Id4 are expressed in the granulosa and theca cells of healthy mammalian ovarian follicles across development. The Ids have two main functions; 1) to maintain proliferation and 2) to inhibit differentiation (4, 6). Such regulation during development ensures that the appropriate level of growth leading to patterning can occur prior to lineage commitment and the terminal differentiation of cells. The adult ovary similarly undergoes persistent tissue remodeling, resulting in the growth and differentiation of somatic follicular cells which house the oocyte. Bi-directional paracrine communication between theca cells, granulosa cells and the oocyte result in the accumulation of genetic and developmental competence for ovulation, fertilization and subsequent embryogenesis. Id proteins are differentially expressed throughout development in many species including the mouse and Xenopus laevis and can have non-redundant actions as well as cell-specific roles (8, 28). We propose a role for the Id proteins in the sheep ovary that might involve transcriptional regulation critical for normal folliculogenesis that may be analogous to those observed during development.

Cumulus cell specific Id1 expression as well as the somatic cell wide Id protein expression was hypothesized to be regulated by activin and/or BMPs, present in the mammalian ovary, which are known to promote growth and differentiation in granulosa and thecal cells of various species (29-31). The steroidogenic cells of the adult ovary are fairly transient and possess similar characteristics to embryonic tissues or progenitor cells of which there are widespread reports of TGFβ-mediated Id mRNA and protein regulation (15, 16, 18). Here, we show that activin and BMPs can alter Id gene expression in ovine granulosa cells in vitro, and moreover, follicles have the cellular machinery for TGFβ signaling in vivo. This was demonstrated in ovine granulosa and thecal cells by the expression of p-smad 1/5/8 and p-smad 2/3 and the presence of smad 6 and smad 7 inhibitors indicating regulation of TGFβ signaling.

Id proteins may be regulated in ovine granulosa cells by various BMPs and activin that act in an opposing way to balance growth and differentiation appropriately. Id1 and Id3 expression was significantly increased following BMP6 stimulation of granulosa cells in vitro whereas activin A down-regulated these genes. Therefore Id protein regulation in the sheep ovary may be a target not only of BMPs but also activin. The effect of activin on Id function is not well described however our findings are consistent with a previous study reporting a negative association with activin treatment and Id gene expression (17). Reports of the specific expression of BMPs in the ruminant follicle vary (30, 32, 33), however it is probable that BMP4, 6 and 7 have actions in these cells and may all regulate Id expression although this is yet to be established. We also found that treating cells with both activin A and BMP6 reduced the BMP-mediated up-regulation of Id genes, suggesting a competing regulatory role for these paracrine molecules.

The universally increased Id1 protein expression observed in peri-oocytic cumulus cells in follicles throughout development suggests a specific regulatory mechanism exerted by Id1 that is likely mediated by paracrine signaling from the oocyte. Although this study revealed TGFβ-mediated signaling via smad 1/5/8 and/or smad 2/3 across the whole follicle, neither pathway could be directly linked to the cumulus-specific Id1 up-regulation. P-smad 2/3 was strongly but not differentially localized in granulosa cells and whilst p-smad 1/5/8 expression was more limited to certain cells this was not specific to the cumulus layer. In fact, co-immunofluorescence established that in the cumulus cells Id1 and p-smad 1/5/8 were not present in the same cell and thus led to a hypothesis that smad 1/5/8 and/or smad 2/3 may indirectly regulate Id1 in these cells. Intermediary factors might include the inhibitory smads and it was observed that smad 6 displayed a more differential expression pattern with some up-regulation in the cumulus cells as well as some co-localization with the Id1 protein. Smad 6 may repress smad signaling in some of these cells although the factor(s) regulating such an interaction are unknown and further work is required to investigate this. Specific oocyte-secreted factors in the sheep include TGFβ signaling molecules such as BMP6 (30) and BMP15 (GDF9b) (34) that activate smad 1/5/8 signaling in granulosa cells (35). BMP15 stimulates granulosa cell proliferation whilst inhibiting differentiation (36, 37) and is essential for folliculogenesis in the sheep (34).

Alternatively the lack of co-localization observed between Id1 and p-smad 1/5/8 raises the possibility that the smad 2/3 pathway could in fact be responsible for the up-regulation of Id1 in peri-oocytic cumulus cells. Activin, an activator of smad 2/3, led to a decrease in Id1 message in granulosa cell culture experiments. However, this effect could be cell specific since the majority of granulosa cells in these cultures are mural and the response may well be different in cumulus granulosa cells modulated by factor(s) secreted by the oocyte. GDF-9, an oocyte-secreted TGFβ member closely related to BMP15 (38), is an excellent candidate for triggering such a response. Its signaling pathway is less clear as it can influence both smad 1/5/8 and smad 2/3 responses (39). GDF-9 signaling has been shown to be essential for cumulus cell function where activation of smad 2/3 brings about the up-regulation of cumulus-specific genes (40). This paracrine signaling through smad 2/3 is crucial for normal cumulus cell expansion, resulting in a local milieu enabling the acquisition of oocyte developmental competence (41-43). We showed in vitro that GDF-9 (NS) and BMP15 can in fact increase Id1 gene expression in cultured ovine granulosa cells and both ligands together enhance this response further although we re-iterate that these findings are limited to a largely mural granulosa cell population which may differ from cumulus cells. In context this latter finding is interesting since the synergy between these growth factors is known to be important for granulosa cell proliferation in the sheep (44). We hypothesize that whilst the surrounding granulosa cells produce activin that might negate Id1 expression via smad 2/3, TGFβ signal from the oocyte such as BMP6, BMP15 and/or GDF-9 may lead to Id1 up-regulation in these cumulus cells.

We also demonstrate the down-regulation of Id proteins in the thecal cells of atretic ovine follicles which are not altered in the granulosa cells. Follicular atresia occurs when FSH levels decrease during the follicular phase of the estrous cycle leading to apoptosis in subordinate follicles (45). The present study identifies a differential change in Id protein distribution in these follicles that may be triggered by autocrine or intracrine thecal signaling, which is potentially inhibitory and does not effect granulosa cell expression. As most changes associated with atresia are detected in the granulosa cell population these theca-specific changes are of interest. However theca-specific Id protein down-regulation in atretic follicles is merely associative with no evidence at this stage of a causal relationship with further work required to clarify the significance of such findings.

This study is the first to our knowledge, to carry out a comprehensive immunohistochemical examination of the Id proteins in the adult mammalian ovary. Id1, Id2, Id3 and Id4 are present in the ovine ovarian follicle and may be regulated by BMPs and/or activin via smad signaling. Further studies are required to establish how important the Ids are for the regulation of proliferation and differentiation of steroidogenic cells occurring during folliculogenesis and the physiological significance for fertility and reproductive diseases.


This is a novel study demonstrating the expression of Id1-4 protein in the adult mammalian ovary in vivo with focus on the signaling pathways that may govern such expression, mediated by ovarian growth factors including the BMPs and activin which we have shown can manipulate Id gene expression in vitro.


The authors acknowledge helpful discussions with Dr Andrew Childs, Professor Richard Anderson and Dr Rachel Dickinson. The expert technical support of Eva Gay and animal husbandry of Joan Docherty and her staff is gratefully appreciated.

Support: This work was funded by the MRC (G0500717) studentship to KH, MRC (G00007.01) to ASM which supports JY, and Dr W.C. Duncan is supported by a Senior Clinical Fellowship from the CSO.)


Disclosure summary: The authors confirm that there are no potential conflicts of interest related to this manuscript.


1. Massari ME, Murre C. Helix-Loop-Helix Proteins: Regulators of Transcription in Eucaryotic Organisms. Mol Cell Biol. 2000;20:429–440. [PMC free article] [PubMed]
2. Murre C, McCaw PS, Vaessin H, Caudy M, Jan LY, Jan YN, Cabrera CV, Buskin JN, Hauschka SD, Lassar AB, Weintraub H, Baltimore D. Interactions between heterologous helix-loop-helix proteins generate complexes that bind specifically to a common DNA sequence. Cell. 1989;58:537–544. [PubMed]
3. O’Toole PJ, Inoue T, Emerson L, Morrison IEG, Mackie AR, Cherry RJ, Norton JD. Id Proteins Negatively Regulate Basic Helix-Loop-Helix Transcription Factor Function by Disrupting Subnuclear Compartmentalization. J Biol Chem. 2003;278:45770–45776. [PubMed]
4. Barone MV, Pepperkok R, Peverali FA, Philipson L. Id proteins control growth induction in mammalian cells. Proc Natl Acad Sci USA. 1994;91:4985–4988. [PubMed]
5. Prabhu S, Ignatova A, Park ST, Sun XH. Regulation of the expression of cyclin-dependent kinase inhibitor p21 by E2A and Id proteins. Mol Cell Biol. 1997;17:5888–5896. [PMC free article] [PubMed]
6. Norton JD, Deed RW, Craggs G, Sablitzky F. Id helix--loop--helix proteins in cell growth and differentiation. Trends Cell Biol. 1998;8:58–65. [PubMed]
7. Riechmann Vs, Van Crüchten I, Sablitzky F. The expression pattern of Id4, a novel dominant negative helix-loop-helix protein, is distinct from Id1, Id2 and Id3. Nucleic Acids Res. 1994;22:749–755. [PMC free article] [PubMed]
8. Liu KJ, Harland RM. Cloning and characterization of Xenopus Id4 reveals differing roles for Id genes. Dev Biol. 2003;264:339–351. [PubMed]
9. Norton JD. ID helix-loop-helix proteins in cell growth, differentiation and tumorigenesis. J Cell Sci. 2000;113:3897–3905. [PubMed]
10. Ruzinova MB, Benezra R. Id proteins in development, cell cycle and cancer. Trends Cell Biol. 2003;13:410–418. [PubMed]
11. Korchynskyi O, ten Dijke P. Identification and Functional Characterization of Distinct Critically Important Bone Morphogenetic Protein-specific Response Elements in the Id1 Promoter. J Biol Chem. 2002;277:4883–4891. [PubMed]
12. Massague J. TGF-beta; SIGNAL TRANSDUCTION. Ann Rev Biochem. 1998;67:753–791. [PubMed]
13. Yue J, Mulder KM. Transforming growth factor-[beta] signal transduction in epithelial cells. Pharmacology & Therapeutics. 2001;91:1–34. [PubMed]
14. Ishisaki A, Yamato K, Hashimoto S, Nakao A, Tamaki K, Nonaka K, ten Dijke P, Sugino H, Nishihara T. Differential Inhibition of Smad6 and Smad7 on Bone Morphogenetic Protein- and Activin-mediated Growth Arrest and Apoptosis in B Cells. J Biol Chem. 1999;274:13637–13642. [PubMed]
15. Peng Y, Kang Q, Luo Q, Jiang W, Si W, Liu BA, Luu HH, Park JK, Li X, Luo J, Montag AG, Haydon RC, He T-C. Inhibitor of DNA Binding/Differentiation Helix-Loop-Helix Proteins Mediate Bone Morphogenetic Protein-induced Osteoblast Differentiation of Mesenchymal Stem Cells. J Biol Chem. 2004;279:32941–32949. [PubMed]
16. Samanta J, Kessler JA. Interactions between ID and OLIG proteins mediate the inhibitory effects of BMP4 on oligodendroglial differentiation. Development. 2004;131:4131–4142. [PubMed]
17. Rotzer D, Krampert M, Sulyok S, Braun S, Stark HJ, Boukamp P, Werner S. Id proteins: Novel targets of activin action, which regulate epidermal homeostasis. Oncogene. 2005;25:2070–2081. [PubMed]
18. Hollnagel A, Oehlmann V, Heymer J, Ruther U, Nordheim A. Id Genes Are Direct Targets of Bone Morphogenetic Protein Induction in Embryonic Stem Cells. J Biol Chem. 1999;274:19838–19845. [PubMed]
19. Hillier SG. Regulatory functions for inhibin and activin in human ovaries. J Endocrinol. 1991;131:171–175. [PubMed]
20. Shimasaki S, Zachow RJ, Li D, Kim H, Iemura S-i, Ueno N, Sampath K, Chang RJ, Erickson GF. A functional bone morphogenetic protein system in the ovary. Proceedings of the National Academy of Sciences of the United States of America. 1999;96:7282–7287. [PubMed]
21. Miyoshi T, Otsuka F, Inagaki K, Otani H, Takeda M, Suzuki J, Goto J, Ogura T, Makino H. Differential Regulation of Steroidogenesis by Bone Morphogenetic Proteins in Granulosa Cells: Involvement of Extracellularly Regulated Kinase Signaling and Oocyte Actions in Follicle-Stimulating Hormone-Induced Estrogen Production. Endocrinology. 2007;148:337–345. [PubMed]
22. Johnson AL, Haugen MJ, Woods DC. Role for Inhibitor of Differentiation/Deoxyribonucleic Acid-Binding (Id) Proteins in Granulosa Cell Differentiation. Endocrinology. 2008;149:3187–3195. [PubMed]
23. Johnson AL, Woods DC. Dynamics of avian ovarian follicle development: Cellular mechanisms of granulosa cell differentiation. General and Comparative Endocrinology. 2009;163:12–17. [PubMed]
24. Nicol L, Faure MO, McNeilly JR, Fontaine J, Taragnat C, McNeilly AS. Bone morphogenetic protein-4 interacts with activin and GnRH to modulate gonadotrophin secretion in LbetaT2 gonadotrophs. J Endocrinol. 2008;196:497–507. [PubMed]
25. Campbell BK, Scaramuzzi RJ, Webb R. Induction and maintenance of oestradiol and immunoreactive inhibin production with FSH by ovine granulosa cells cultured in serum-free media. J Reprod Fertil. 1996;106:7–16. [PubMed]
26. Thomas FH, Armstrong DG, Telfer EE. Activin promotes oocyte development in ovine preantral follicles in vitro. Reprod Biol Endocrinol. 2003;1:76. [PMC free article] [PubMed]
27. Young JM, Juengel JL, Dodds KG, Laird M, Dearden PK, McNeilly AS, McNatty KP, Wilson T. The activin receptor-like kinase 6 Booroola mutation enhances suppressive effects of bone morphogenetic protein 2 (BMP2), BMP4, BMP6 and growth and differentiation factor-9 on FSH release from ovine primary pituitary cell cultures. J Endocrinol. 2008;196:251–261. [PubMed]
28. Benezra R, Davis RL, Lockshon D, Turner DL, Weintraub H. The protein Id: A negative regulator of helix-loop-helix DNA binding proteins. Cell. 1990;61:49–59. [PubMed]
29. Knight PG, Glister C. Local roles of TGF-[beta] superfamily members in the control of ovarian follicle development. Animal Reproduction Science. 2003;78:165–183. [PubMed]
30. Juengel JL, Reader KL, Bibby AH, Lun S, Ross I, Haydon LJ, McNatty KP. The role of bone morphogenetic proteins 2, 4, 6 and 7 during ovarian follicular development in sheep: contrast to rat. Reproduction. 2006;131:501–513. [PubMed]
31. Zhao J, Taverne MAM, van der Weijden GC, Bevers MM, van den Hurk R. Effect of Activin A on In Vitro Development of Rat Preantral Follicles and Localization of Activin A and Activin Receptor II. Biol Reprod. 2001;65:967–977. [PubMed]
32. Glister C, Kemp CF, Knight PG. Bone morphogenetic protein (BMP) ligands and receptors in bovine ovarian follicle cells: actions of BMP-4, −6 and −7 on granulosa cells and differential modulation of Smad-1 phosphorylation by follistatin. Reproduction. 2004;127:239–254. [PubMed]
33. Campbell BK, Souza CJH, Skinner AJ, Webb R, Baird DT. Enhanced Response of Granulosa and Theca Cells from Sheep Carriers of the FecB Mutation in Vitro to Gonadotropins and Bone Morphogenic Protein-2, −4, and −6. Endocrinology. 2006;147:1608–1620. [PubMed]
34. Galloway SM, McNatty KP, Cambridge LM, Laitinen MP, Juengel JL, Jokiranta TS, McLaren RJ, Luiro K, Dodds KG, Montgomery GW, Beattie AE, Davis GH, Ritvos O. Mutations in an oocyte-derived growth factor gene (BMP15) cause increased ovulation rate and infertility in a dosage-sensitive manner. Nat Genet. 2000;25:279–283. [PubMed]
35. Moore RK, Otsuka F, Shimasaki S. Molecular basis of bone morphogenetic protein-15 signaling in granulosa cells. J Biol Chem. 2002:M207362200. [PubMed]
36. Otsuka F, Yao Z, Lee T-h, Yamamoto S, Erickson GF, Shimasaki S. Bone Morphogenetic Protein-15. IDENTIFICATION OF TARGET CELLS AND BIOLOGICAL FUNCTIONS. J Biol Chem. 2000;275:39523–39528. [PubMed]
37. Otsuka F, Yamamoto S, Erickson GF, Shimasaki S. Bone Morphogenetic Protein-15 Inhibits Follicle-stimulating Hormone (FSH) Action by Suppressing FSH Receptor Expression. J Biol Chem. 2001;276:11387–11392. [PubMed]
38. Dube JL, Wang P, Elvin J, Lyons KM, Celeste AJ, Matzuk MM. The Bone Morphogenetic Protein 15 Gene Is X-Linked and Expressed in Oocytes. Mol Endocrinol. 1998;12:1809–1817. [PubMed]
39. Mazerbourg S, Klein C, Roh J, Kaivo-Oja N, Mottershead DG, Korchynskyi O, Ritvos O, Hsueh AJW. Growth Differentiation Factor-9 Signaling Is Mediated by the Type I Receptor, Activin Receptor-Like Kinase 5. Mol Endocrinol. 2004;18:653–665. [PubMed]
40. Diaz FJ, Wigglesworth K, Eppig JJ. Oocytes determine cumulus cell lineage in mouse ovarian follicles. J Cell Sci. 2007;120:1330–1340. [PubMed]
41. Gilchrist RB, Ritter LJ, Armstrong DT. Oocyte–somatic cell interactions during follicle development in mammals. Anim Reprod Sci. 2004;82-83:431–446. [PubMed]
42. Dragovic RA, Ritter LJ, Schulz SJ, Amato F, Thompson JG, Armstrong DT, Gilchrist RB. Oocyte-Secreted Factor Activation of SMAD 2/3 Signaling Enables Initiation of Mouse Cumulus Cell Expansion. Biol Reprod. 2007;76:848–857. [PubMed]
43. Li Q, Pangas SA, Jorgez CJ, Graff JM, Weinstein M, Matzuk MM. Redundant Roles of SMAD2 and SMAD3 in Ovarian Granulosa Cells In Vivo. Mol Cell Biol. 2008;28:7001–7011. [PMC free article] [PubMed]
44. McNatty KP, Juengel JL, Reader KL, Lun S, Myllymaa S, Lawrence SB, Western A, Meerasahib MF, Mottershead DG, Groome NP, Ritvos O, Laitinen MPE. Bone morphogenetic protein 15 and growth differentiation factor 9 co-operate to regulate granulosa cell function. Reproduction. 2005;129:473–480. [PubMed]
45. Hussein MR. Apoptosis in the ovary: molecular mechanisms. Hum Reprod Update. 2005;11:162–178. [PubMed]