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. 2010 Mar 10;151(5):2319–2330. doi: 10.1210/en.2009-1489

Expression of E-Cadherin and N-Cadherin in Perinatal Hamster Ovary: Possible Involvement in Primordial Follicle Formation and Regulation by Follicle-Stimulating Hormone

Cheng Wang 1, Shyamal K Roy 1
PMCID: PMC2869259  PMID: 20219978

Abstract

We examined the expression and hormonal regulation of E-cadherin (CDH1) and N-cadherin (CDH2) with respect to primordial follicle formation. Hamster Cdh1 and Cdh2 cDNA and amino acid sequences were more than 90% similar to those of the mouse, rat, and human. Although CDH1 expression remained exclusively in the oocytes during neonatal ovary development, CDH2 expression shifted from the oocytes to granulosa cells of primordial follicles on postnatal day (P)8. Subsequently, strong CDH2 expression was restricted to granulosa cells of growing follicles. Cdh2 mRNA levels in the ovary decreased from embryonic d 13 through P10 with a transient increase on P7, which was the day before the appearance of primordial follicles. Cdh1 mRNA levels decreased from embryonic d 13 through P3 and then showed a transient increase on P8, coinciding with the formation of primordial follicles. CDH1 and CDH2 expression were consistent with that of mRNA. Neutralization of FSH in utero impaired primordial follicle formation with an associated decrease in Cdh2 mRNA and CDH2, but an increase in Cdh1 mRNA and CDH1 expression. The altered expression was reversed by equine chorionic gonadotropin treatment on P1. Whereas a CDH2 antibody significantly reduced the formation of primordial and primary follicles in vitro, a CDH1 antibody had the opposite effect. This is the first evidence to suggest that primordial follicle formation requires a differential spatiotemporal expression and action of CDH1 and CDH2. Further, FSH regulation of primordial follicle formation may involve the action of CDH1 and CDH2.

Primordial follicle formation is associated with differential expression and action of cadherins, and the process is regulated by FSH.

Alterations in the assembly or disassembly of adherent junctions play a major role in the morphogenetic processes of many organs (1). The dynamic physiological regulation of the adhesive functions of classic cadherins, a family of transmembrane glycoproteins, at the cell surface is especially important for morphogenesis (2). Cdh1 and Cdh2 are expressed in rat (3,4), mouse (5,6), pig (7), and human (8) ovarian cells; however, the pattern of relative localization in ovarian cells remains inconsistent across species. Estrogen affects Cdh1 and Cdh2 mRNA levels in the mouse ovary (9) and rat granulosa cells (3,10), but whether cadherins are important for primordial follicle formation warrants thorough investigation. E-cadherin (CDH1) seems to be involved in establishing the germ cell lineage (11), oocyte growth, and the acquisition of meiotic competence during gonad development in mice (12). On the other hand, N-cadherin (CDH2)-mediated adhesion of rat granulosa cells in culture prevents apoptosis (13,14,15). These lines of evidence suggest that CDH1 and CDH2 play an important role in folliculogenesis, but whether these two cadherins are differentially expressed in neonatal ovary cells during somatic cell and oocyte assembly forming primordial follicles, and the biological relevance of such expression, remains virtually unknown.

A family of intracellular proteins that bind to CDH1 and CDH2 is catenin. The mostly studied members are α-catenin, β-catenin (CTNNB1), and γ-catenin (JUP) (2). Of the three, CTNNB1 binds to the CDH1 and CDH2 at the plasma membrane and to the CTNNA at the N-terminal site, which connects to the β-actin and several other actin-binding proteins. There is also a cytoplasmic form of CTNNB1, which serves as an intracellular signal transduction molecule to mediate Wnt signaling (1,2). Cytoplasmic CTNNB1 plays a significant role in tissue morphogenesis and carcinogenesis (16). Similar to CTNNB1, JUP can also bind to CDH1 and CDH2 at the CTNNB1-binding region; however, it does not participate in Wnt signaling. Although CTNNB1 have been studied in the context of cancer cell adhesion and migration and Wnt signaling, the types of catenin associated with cadherins in ovarian cells, especially during perinatal follicular morphogenesis, remains unknown.

The first cohort of primordial follicles appears in the hamster ovary in vivo on postnatal day (P)8 (17) and in vitro after 9 d of culture of embryonic day (E)15 ovaries (18). This unique development offers an extended postnatal window to study the role of cadherins in primordial follicle formation. We have shown that FSH (17), as well as estradiol-17β (19,20), facilitate somatic cell and oocyte assembly leading to the formation of primordial follicles; however, whether the underlying mechanism involves CDH1 and CDH2 remains unknown. The aim of the present study was to examine the FSH regulation and biological significance of Cdh1 mRNA, CDH1, Cdh2 mRNA, and CDH2 expression in ovary cells during perinatal ovarian morphogenesis with respect to the formation of primordial follicles.

Materials and Methods

Chemicals and animals

The mouse monoclonal antibodies to CDH2 and CDH1 were purchased from BD Transduction Laboratory (San Jose, CA). A rabbit polyclonal FSH antiserum that neutralized hamster FSH was prepared in the laboratory and tested for its efficiency to block primordial follicle development in the hamster (17,21). CDH1 neutralization antibody was purchased from Takara Bio, Inc. (Otsu, Japan), CDH2 neutralization antibody was purchased from Sigma Chemical Co. (St. Louis, MO). Second antibody conjugated to Alexa 488 was purchased from Invitrogen (Carlsbad, CA), equine chorionic gonadotropin (eCG) (2000 IU/mg solid) was purchased from Sigma Chemical Co., and PCR chemicals were from Roche Molecular Biochemicals (Indianapolis, IN), Pharmacia Biotech Boehringer (Piscataway, NJ), and Invitrogen. Quantitative (q)PCR primers and probes were synthesized in the Eppley DNA Synthesis Core Facility (University of Nebraska Medical Center). Second antibodies for Western blotting chemiluminescence were from Jackson ImmunoResearch, Inc. (West Grove, PA), enhanced chemiluminescence advance Western blot detection kit was from GE Healthcare (Buckinghamshire, UK), Optitran nitrocellulose transfer membrane was from Schleicher & Schuell Bioscience (Dassel, Germany), the RNeasy mini kit was from QIAGEN, Inc. (Valencia, CA), and DMEM and other cell culture media were from Invitrogen. All other molecular-grade chemicals were purchased from Sigma, Fisher (Pittsburgh, PA), or United States Biochemical (Cleveland, OH).

Golden hamsters (90–100 g) were purchased from Charles River Laboratories (Charles River, MA) and maintained in a climate-controlled room with 14-h light, 10-h dark cycle with free access to food and water. The use of hamsters for this study was according to the Institutional Animal Care and Use Committee and the United States Department of Agriculture guidelines and was approved by the Institutional Animal Care and Use Committee. Females with at least three consecutive estrous cycles were mated with males in the evening of proestrus, and the presence of sperm in the vagina the next morning was considered the first day of pregnancy. Hamster gestation lasts for 16 d, and pups are born on the 16th day of gestation, which we considered the first day of postnatal life (P1).

Partial cloning of hamster Cdh1 mRNA and Cdh2 mRNA

Hamster Cdh1 and Cdh2 cDNA were cloned using RT and PCR amplification of the cDNA. cDNA was amplified from total RNA of P8 ovaries using previously described protocols (22,23,24). The primer sequences used for amplifying and quantifying hamster Cdh1 and Cdh2 cDNA were presented in Supplemental Table 1 published on The Endocrine Society’s Journals Online web site at http://endo.endojournals.org.

The primers were designed based on human (NM_001792), rat (NM_031333), and mouse (BC022107) Cdh1 and Cdh2 and hamster ActB (AJ312092) cDNA using Vector NTI software (Invitrogen) and Primer3 software (25). For the cloning, the PCR conditions were as follows: 15 min at 95 C, then 35 cycles of 1 min at 94 C, 1 min at 55 C, and 1 min at 72 C, followed by 10 min at 72 C. cDNA was cloned into a PCR4-TOPO4 plasmid vector (Invitrogen) and sequenced in the DNA sequencing core facility (University of Nebraska at Lincoln, Lincoln, NE, and McLab, San Francisco, CA). The sequences were identified as Cdh1 and Cdh2 and ActB by BLASTing [National Center for Biotechnology Information (NCBI), Bethesda, MD]. The similarity between the hamster Cdh1 and Cdh2 cDNA sequences with the corresponding stretch of rat, mouse, and human Cdh1 and Cdh2 cDNA sequences were determined by alignment using the Vector NTI software. Partial amino acid sequences were deduced from the cDNA, and all sequences were deposited in the NCBI GenBank.

Expression of Cdh1 and Cdh2 mRNA and CDH1 and CDH2 and CTNNB1 during perinatal development with respect to the formation of primordial follicles

Ovaries were collected from E13 through E15 and P1 through P15, and 6-μm-thick frozen sections were used for immunofluorescence detection of CDH1 and CDH2 and CTNNB1. A second set of frozen ovaries for each day of development was used for Western blot determination of the relative levels of cadherin proteins. A third set of ovaries for each day of development was collected for RNA preparation, which was used for qRT-PCR determination of the levels of Cdh1, Cdh2, and ActB mRNA.

The association between CTNNB1 and JUP with CDH1 or CDH2 was also examined by immunoprecipitating the protein complex from P8 hamster ovary homogenates with the antibody specific to CTNNB1 or JUP and by immunoblot detection of the type of cadherin associated with specific types of catenin.

Effect of anti-FSH serum on ovarian CDH1 and CDH2 mRNA and CDH1 and CDH2

Pregnant hamsters were injected sc with 200 μl of an anti-FSH serum on 12th day of gestation as described previously (17). After birth, one set of pups received a sc injection of 0.5% BSA in saline (vehicle for eCG), whereas the other received 20 IU eCG on P1. Ovaries were collected on P8 for quantification of Cdh1 and Cdh2 mRNA and immunofluorescence detection of CDH1 and CDH2. Equine CG was used because of its longer half-life in vivo, thus avoiding repeated injections to newborn pups and its ability to stimulate follicular development in the hamster (26,27,28). Moreover, eCG did not cross-react with the FSH antiserum; hence, could bypass the antibody neutralization. Although the FSH-receptor mRNA was expressed in the hamster ovary from E13 and onwards, LH receptor mRNA did not appear in postnatal hamster ovarian cells until P13 (Roy, S. K., unpublished observation).

Effect of CDH1 or CDH2 neutralization antibodies on the formation and development of primordial follicles in neonatal hamster ovary

Antibodies against the functional-binding motifs of CDH1 and CDH2 were used for blocking the CDH1 and CDH2 action in in vitro ovary culture system. E15 ovaries were cultured for 9 d in DMEM supplemented with 0.1 mg/ml human recombinant insulin, 6.25 μg/ml human transferrin, 6.25 ng/ml seleneous acid, 5 mg/ml BSA, and 5.35 ng/ml linoleic acid under 5% CO2 in air as described previously (18,29) in the presence of 50 μg/ml CDH1 neutralizing antibody (30,31) or 20 μg/ml CDH2 neutralization antibody (32,33). Control culture received equal amounts of normal mouse IgG. The medium was replaced with or without the antibody every 48 h. Ovaries were retrieved, fixed in Bouin’s fixative, and the percentage of primordial follicle formation was quantified by morphometry as described previously (17,29,34). In an initial experiment, the complete penetration of IgG in the ovary during culture was ensured using 20 μg/ml Alexa 488 conjugated mouse IgG and confocal microscopy.

Quantitative RT-PCR of Cdh1 and Cdh2 mRNA

Ovarian RNA was isolated using the RNeasy mini kit, quantified by Ribogreen kit (Invitrogen), and the levels of Cdh1, Cdh2, and ActB mRNA were determined by RT-qPCR using respective cRNA as standards as described previously for other gene transcripts (24,29). Quantitative PCR primers and probes were designed based on hamster Cdh1, Cdh2, and ActB cDNA sequences using the Primer Express software (Applied Biosciences, Foster City, CA). The authenticity of the qPCR products was verified by nucleic acid sequencing and by withholding reverse transcriptase from the reaction. The values were presented as ratios of cadherin and actin mRNA.

Immunoblot determination of CDH1 or CDH2 during perinatal ovary development and immunoprecipitation of cadherin and catenin complex from P8 hamster ovaries

Ovaries from E13 through P15 were sonicated in lysis buffer (10 mm Tris-HCl, pH 7.4; 100 mm NaCl; 1 mm EDTA; 1 mm EGTA; 1 mm NaF; 20 mm Na4P2O7; 1% Triton X-100; 10% glycerol; 0.1% sodium dodecyl sulfate; and 0.5% deoxycholate) containing a protease inhibitor cocktail and phenylmethylsulfonylfluoride on ice. After measuring the protein concentration in 14,000 × g supernatants by Micro BCA protein assay kit (Pierce, Rockford, IL), 20-μg protein was fractionated in a SDS-PAGE, electrotransferred to an Optitran nitrocellulose membrane, and probed with the anti-CDH1, anti-CDH2, or anti-β-tubulin (TUBB) antibody and an appropriate second antibody. The chemiluminescence signal was developed using the Advance Western blot detection kit (GE Healthcare), and recorded by a UVP gel documentation system (UVP, Upland, CA). The data were normalized with tubulin and presented as a ratio. Each group had at least three replicates of samples collected from three different animals.

CTNNB1 or JUP was immunoprecipitated overnight at 4 C from 200 μg of P8 hamster ovary protein in PBS (pH 7.4) containing 0.1% Triton X-100 and a protease inhibitor cocktail using the antibody specific to CTNNB1 or JUP, and protein A/G-agarose as described previously (35). The protein and bead complex was washed four times in ice-cold PBS containing 0.1% Triton X-100, and the immunoprecipitated protein complex was extracted using 3× sodium dodecyl sulfate buffer containing β-mercaptoehtanol, denatured in a boiling water bath for 5 min, resolved in 10% polyacrylamide gel, transferred to the Optitran membrane, and probed simultaneously with either CDH1 and JUP or CDH2 and CTNNB1 specific antibodies using previously described protocols (35,36).

Immunofluorescence localization of CDH1 or CDH2

Frozen sections at 6 μm were fixed in freshly prepared ice-cold 4% paraformaldehyde in PBS (pH 7.4) for 10 min and stained for CDH1 and CDH2 protein as described previously (24). The images were captured by a QImaging digital camera (QImaging, Surrey, British Columbia, Canada) and Openlab (Perkin-Elmer, Waltham, MA) image analysis software. The exposure time of the camera was set for subtracting background fluorescence that was present in sections incubated with the nonimmune IgG of the host species. CDH1 or CDH2 specific fluorescence immunosignal was merged with the nuclear signal (4′,6-diamidino-2-phenylindole) to determine the cellular site of protein expression. The nuclear signal was blue, whereas the CDH1 and CDH2 signals were green.

To determine whether CDH1 or CDH2 in developing hamster ovarian cells was coupled with CTNNB1 or JUP, CDH1 or CDH2 was costained with CTNNB1 or JUP using a previously described protocol (24).

We examined whether CDH1 expression would turn off in the oocytes, but remain expressed in juxtaposed granulosa cells after the formation of follicles by staining the zona pellucida with wheat germ agglutinin (WGA)-Alexa 594. The objective was to clearly identify the boundaries of the oocyte plasma membrane and granulosa cells. Sections of P20 ovaries with the full complement of preantral follicle development were stained with CDH2 as described earlier, but WGA-Alexa 594 exposure was limited to 30 min at room temperature to avoid over staining.

We examined whether the expression of CDH2 in somatic cells depended on their juxtaposition with the oocytes by staining the oocytes in P8 and P9 ovarian sections with the mouse vasa homolog (MVH) antibody. Further, to determine whether somatic cells apposed to the oocytes were indeed granulosa cells committed to form follicles, sections of P9 ovaries were costained for the presence of laminin, which is a component of the basal lamina surrounding the follicles.

Effects of neutralization of CDH1 or CDH2 on primordial follicle formation in vitro

E15 ovaries were cultured in vitro for 9 or 10 d as described previously (17,29) in the absence or presence of 20 μg/ml normal mouse IgG-Alexa 488, 20 or 50 μg/ml normal mouse IgG, 50 μg/ml mouse monoclonal anti-CDH1, or 20 μg/ml anti-CDH2 antibody. Fresh medium with IgG or antibody was replaced every 48 h. Ovaries were retrieved after either 9 d (for CDH1 antibody) or 10 d (for CDH2 antibody) of culture and examined for the distribution of fluorescence-labeled IgG or processed for morphological assessment of primordial follicle formation as described previously (17,19,20).

Statistics

All cultures and immunofluorescence localizations were repeated at least three times using ovaries from different fetuses. Ovaries from untreated, antiserum-treated, or eCG-treated groups were cultured in parallel. Fetal ovaries from each pregnant hamster were pooled to obtain one sample for each fetal age. There were three pregnant hamsters for each fetal age to obtain three replicates. RNA and protein samples for each P were prepared from ovaries pooled from three litters, and there were at least three samples for each P. All quantitative data were analyzed by one-way ANOVA with Scheffé’s post hoc test using StatView software (SAS Institute, Inc., Cary, NC). The level of significance was 5%.

Results

Cloning and characterization of hamster Cdh1 and Cdh2 cDNAs

The sequences of hamster Cdh1 and Cdh2 cDNA were submitted to the GenBank (NCBI accession nos. DQ237897 and DQ237892). The 1796-bp hamster Cdh2 cDNA was 93.8, 92.0, and 87.8% similar to the corresponding reading frame of mouse, rat, and human cDNA, respectively. The deduced amino acid sequence was 97.2, 97.2, and 95.2% similar to that of mouse, rat, and human, respectively. The 1100-bp hamster Cdh1 cDNA was 97.5, 94.3, and 92.2% similar to the corresponding reading frame of mouse, rat, and human cDNA, respectively, whereas the deduced amino acid sequence was 98.1, 97.9, and 95.9% similar to that of mouse, rat, and human, respectively. The feature sequences of the CDH2, such as the repeat calcium-binding motifs, Asp-×-Asn-Asp-Asn, Asp-×-Asp, and Leu-Asp-Asn-Glu, and the classical cadherin adhesion recognition sequence, His-Ala-Val and Ile-Asn-Pro-Ile-Ser-Gly-Gln in the extracellular domain 1 (37), were present in the deduced amino acid sequence of the hamster CDH2.

Developmental expression of Cdh1 and Cdh2 mRNA and proteins in perinatal hamster ovaries

The rationale was to examine whether the developmental expression of Cdh1, Cdh2, CDH1, and CDH2 were temporally correlated with the formation and development of primordial follicles. Cdh2 mRNA levels were proportionately much higher than that of Cdh1, suggesting that Cdh2 was the major form of cadherin in neonatal hamster ovaries. Cdh1 mRNA levels decreased significantly from E14 through P6 (Fig. 1A). The mRNA levels declined further on P7 and remained low through P12 with a transient increase on P8 (Fig. 1A). Cdh2 mRNA levels decreased significantly on E14 and remained through P5 (Fig. 1A). The Cdh2 mRNA decreased further on P6 followed by a small, but significant increase on P7, and then remained steady through P12 (Fig. 1A). Tubb mRNA levels did not show significant fluctuations throughout the perinatal period (data not shown).

Figure 1.

Figure 1

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Expression of Cdh1 and Cdh2 mRNA (A) and CDH1 and CDH2 (B) in perinatal hamster ovary. Cdh1 and Cdh2 mRNA and protein levels were presented as ratios of Act and TUBB, respectively. Each bar represented a mean mRNA ± sem. Bars with same letter are not significantly (P < 0.05) different from each other.

Consistent with the pattern of mRNA expression, CDH1 protein levels decreased through P5 followed by a gradual increase by P7 (Fig. 1B). CDH1 protein levels dropped transiently on P8 after a drop in mRNA levels on P7, coinciding with the appearance of primordial follicles (Fig. 1B). CDH2 protein decreased through P3 and then remained steady through P12 (Fig. 1B).

Localization of CDH1 and CDH2 in developing hamster ovarian cells

Images representing changes in the spatiotemporal expression of CDH1 and CDH2 proteins in the context of follicle formation were furnished. MVH (red) expression identified the oocytes in all sections. CDH1 (green) was present in the fetal hamster oocytes as early as E13 (data not shown), and by E14, oocytes in the oocyte clusters (egg nest) showed notable expression of CDH1, whereas undifferentiated somatic cells had none (Fig. 2A). Strong CDH1 expression remained in the oocytes by P7, but somatic cells surrounding the egg nest had no expression (Fig. 2B). With the appearance of primordial follicles on P8, CDH1 remained constraint in the oocytes, primarily in the oocyte plasma membrane and cytoplasm (Fig. 2C, inset), but granulosa cells of primordial follicles or interstitial cells had no appreciable expression (Fig. 2C). This unique expression pattern was more prominent on P9 (Fig. 2D) and P12 (Fig. 2E), when ovaries contained many primary and early secondary follicles. Interestingly, neutralization of FSH in utero by an FSH antiserum administered on the gestation d 12, which interfered with primordial follicle formation on P8 (17), resulted in marked up-regulation of CDH1 expression on P8 not only in the oocytes, but also in somatic cells throughout the ovary (Fig. 2F), and the effect was completely reversed by eCG administered on P1 (Fig. 2G). Previously, we showed that eCG stimulated primordial and primary follicle formation in antiserum-treated hamsters (17).

Figure 2.

Figure 2

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Localization of CDH1 in hamster ovaries during normal perinatal development and after in utero exposure to an FSH antiserum. A, E14; B, P7; C, P8; D, P9; E, P12. F and G, P8 ovaries from hamsters exposed in utero to an FSH antiserum on gestation d 12 and treated on P1 (F) with saline or (G) 20 IU eCG. O, Oocytes; OC, oocyte clusters or egg nests; S, somatic cells; S0, primordial follicles; S1, primary follicles; S3, stage secondary follicles; GC, granulosa cells; IC, interstitial cells. Green, CDH1; red, MVH; blue, nuclei. Scale bar, 10 μm. C (inset), Higher magnification of a primordial follicle.

CDH2 expression in the ovary also occurred as early as E13 (data not shown); however, by E14, CDH2 immunosignal was located primarily in the oocytes in the clusters; but in contrast to CDH1, the expression was also present in somatic cells (Fig. 3A). By P6, CDH2 expression was located in the apical side of somatic cells surrounding the egg nest, whereas oocytes themselves did not have noticeable expression (Fig. 3B). With the formation of primordial follicles by P8, CDH2 expression was strictly associated with the newly formed granulosa cells or somatic cells near the egg clusters (Fig. 3C). By P9, granulosa cells of primary and secondary follicles (Fig. 3D, inset) showed exclusive expression of CDH2, whereas interstitial cells or oocytes had none (Fig. 3D). CDH2 expression in the granulosa cells became more prominent by P12 when large secondary follicles appeared (Fig. 3E). In contrast to CDH1, neutralization of FSH by the FSH antiserum in utero resulted in marked suppression of CDH2 expression in somatic cells on P8, although no appreciable change was noted for MVH expression (Fig. 3F). Equine CG treatment on P1 of FSH antiserum-treated hamsters restored CDH2 expression in the granulosa cells, but not in the interstitial cells (Fig. 3G).

Figure 3.

Figure 3

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Localization of CDH2 in hamster ovaries during normal perinatal development and after in utero exposure to an FSH antiserum. A, E14; B, P6; C, P8; D, P9; E, P12. F and G, P8 ovaries from hamsters exposed in utero to an FSH antiserum on gestation d 12 and treated on P1 (F) with saline or (G) 20 IU eCG. O, Oocytes; OC, oocyte clusters or egg nests; S, somatic cells; S0, primordial follicles; S1, primary follicles; S3, stage 3 secondary follicles; GC, granulosa cells; IC, interstitial cells. Green, CDH2; red, MVH; blue, nuclei. Scale bar, 10 μm. D (inset), Primary follicle.

Further examination revealed that CDH1 was expressed primarily in the oocytes on E15, whereas occasional granulosa cells adjacent to the egg nest showed modest expression (Fig. 4A). In contrast, granulosa cells of fully formed follicles in P10 ovaries expressed no CDH1, which was strictly restricted to the oocytes (Fig. 4B). CDH1 protein was located in the oocyte plasma membrane as well as in the cytoplasm, indicating that CDH1 may play an important role in oocyte biology. CDH2 was located in the oocytes and in somatic cells surrounding the egg nest on E15 (Fig. 4C), but it was located exclusively in the granulosa cells by P10 (Fig. 4D).

Figure 4.

Figure 4

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CDH1 expression in (A) E15 and (B) P10 ovaries and CDH2 expression in (C) E15 and (D) P10 ovaries. Green, CDH1 and CDH2; red, MVH; blue, nuclei. O, Oocytes; S, somatic cells; S1, primary follicles; S3, stage 3 secondary follicles; GC, granulosa cells. Scale bar, 10 μm.

Consistent with cadherin protein expression in the ovaries of FSH antiserum-treated hamsters, Cdh1 mRNA levels increased more than 2-fold, whereas Cdh2 mRNA levels declined almost 4-fold after the neutralization of endogenous FSH (Fig. 5). Equine CG replacement on P1 reversed the effect (Fig. 5).

Figure 5.

Figure 5

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FSH regulation of Cdh1 and Cdh2 mRNA expression in postnatal hamster ovary in vivo. Each bar represented a ratio of cadherins and Act mRNA ± sem. AS, Antiserum. Bars with the same letter are not significantly (P < 0.05) different from each other.

Examination of the granulosa cell location of CDH2

WGA staining clearly and specifically depicted the extent of the zona boundary (Fig. 6A). Although CDH2 was clearly located in the oocyte plasma membrane during early postnatal ovary development (Fig. 4C), the expression was switched off once the oocytes were surrounded by somatic cells committed to form the granulosa cells. CDH2 immunosignal was present exclusively in the granulosa cells, whereas thecal cells had no immunosignal (Fig. 6B). Granulosa cells adjacent to the zona pellucida showed accumulation of CDH2, which extended in the granulosa cell processes traversing the zona pellucida and apposed to the oocyte plasma membrane (Fig. 6B).

Figure 6.

Figure 6

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Expression of CDH2 (A and B) at the interface of granulosa cells and oocyte in follicles of P20 ovaries, and (C and D) during primordial follicle formation in P9 ovaries. A, Zona pellucida stained with WGA-Alexa 594 showing the oocyte border; B, merger of A with the image showing CDH2 expression in the follicle. Punctate CDH2 immunosignal was associated with granulosa cell processes (arrows) traversing the zona pellucida and apposed to the oocyte plasma membrane. Note total absence of CDH2 in thecal cells or in the oocyte. Red, Zona pellucida; green, CDH2; blue, nuclei. C, Laminin expression surrounding the newly formed granulosa cells of primordial follicles in P8 ovaries. D, Merger of C with the image showing CDH2 expression. Note that cells that were part of the follicles expressed laminin and CDH2, whereas cells elsewhere had neither expression. Red, CDH2; green, laminin; white, nuclei. O, Oocytes; GC, granulosa cells; ZP, zona pellucida; S0, primordial follicles; Th, theca; IC, interstitial cells. Scale bar, 10 μm.

CDH2 expression is a hallmark of granulosa cells

We wanted to know whether CDH2 expression in somatic cells occurred when they commit to differentiate into granulosa cells forming primordial follicles. Ovarian follicles are characterized by the presence of a basement membrane (BM) that separates granulosa cell compartment from surrounding cells. Laminin is one of the matrix proteins in the BM and participates in the regulation of cell functions (38,39,40). We reasoned that if somatic cells surrounding the oocyte committed to form pregranulosa cells, then they should express laminin. Distinct laminin immunosignal was detected around each granulosa cell of the primordial follicles forming a BM outline in P9 ovaries; however, interstitial cells immediately apposed to the flattened granulosa cells had no expression (Fig. 6C). The deposition of laminin demarcated the follicle boundary and isolated the follicular structure (Fig. 6C). Interestingly, CDH2 expression also occurred in the granulosa cells of primordial follicles, which were located exclusively within the boundary demarcated by laminin (Fig. 6D).

Colocalization of CDH1 or CDH2 with CTNNB1

The optimal activity of cell-cell communication depends upon intracellular linkage of the CDH1 and CDH2 linking to actin microfilaments at zonula and fascia adherent junctions, respectively. Both CTNNB1 and JUP can bind with CDH2 or CDH1 (41,42); however, the interaction of cadherin and catenin with respect to ovarian follicular development remained unknown. To address this issue, we wanted to determine which of the catenins interacted with CDH2 or CDH1 in developing hamster ovarian cells using immunofluorescence colocalization and immunoprecipitation. Whereas JUP immunosignal was modest (data not shown), CTNNB1 was expressed both in the oocytes and somatic cells as early as E13 (data not shown) and was prominent by E15 (Fig. 7, A and G). CTNNB1 colocalized with CDH1 (Fig. 7B) in the oocytes as well as in somatic cells of E15 ovaries (Fig. 7C). After the formation of primordial follicles on P8, CTNNB1 was localized in the oocytes, granulosa cells, and other cells in the ovary (Fig. 7, D and J). CDH1 was associated only with the oocytes (Fig. 7E) and colocalized with the CTNNB1 (Fig. 7F). CTNNB1 and CDH2 colocalized in the oocytes and somatic cells on E15, but for other cells (Fig. 7I, arrowheads) no such colocalization was apparent. This was more prominent in P8 ovaries, in which CTNNB1 in the interstitial cells (Fig. 7J) did not show any colocalization with CDH2 (Fig. 7K), which was primarily expressed in the granulosa cells (Fig. 7L). The colocalization of CTNNB1 and CDH2 became more evident in multilayered follicles in P15 ovaries (Fig. 7, M–O).

Figure 7.

Figure 7

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Colocalization of CDH1 (A–F) or CDH2 (G–O) with CTNNB1 before and after primordial follicle formation in developing hamster ovaries. A and G, E15 CTNNB1; D and J, P8 CTNNB1; B, E15 CDH1; C, Merged A and B; E, P8 CDH1; F, merged D and F; H, E15 CDH2; I, Merged G and H; K, P8 CDH2; L, Merged J and K; M, P15 CDH2; N, CTNNB1; O, Merged M and N. Green, CTNNB1; red, CDH1 or CDH2; yellow-orange; merged red and green; blue, nuclei. O, Oocytes; S, somatic cells; OC, oocyte clusters; GC, granulosa cells; IC, interstitial cells; S0, primordial follicles; S5, secondary follicles with five layers of GC; Th, theca. Scale bar, 10 μm.

Immunoprecipitation results indicated that although CTNNB1 and JUP were present in P8 hamster ovaries, only CTNNB1 interacted with both CDH1 and CDH2 in vivo (Fig. 8, A and B). The immunohistochemical study did not show any colocalization of JUP with CDH1 or CDH2 (data not shown).

Figure 8.

Figure 8

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A and B, In vivo interaction of CDH1 or CDH2 with CTNNB1 and JUP in vivo in P8 ovaries. P8 ovarian samples were mixed with either mouse monoclonal CTNNB1 or rabbit polyclonal JUP antibody (ab) to immunoprecipitate (IP) cadherins bound to the antigens. After the electrophoretic separation, the blot was probed simultaneously with (A) mouse monoclonal CDH1and JUP or (B) CDH2 and CTNNB1 antibodies due to appreciable difference in the molecular weight of the two proteins. Lane 1, Whole ovary extract without the immunoprecipitation (IP); lane 2, ovary homogenate immunoprecipitated with the CTNNB1 antibody; and lane 3, ovary homogenate immunoprecipitated with the JUP antibody. Arrows in A indicate the migration of CDH1 and JUP; arrows in B indicate the migration of CDH2 and CTNNB1. Effect of CDH1 neutralization antibody GC4 (C) or CDH2 neutralization antibody ECCD1 (D) on the formation and development of primordial follicles in vitro. E15 ovaries were cultured for 9 d with normal mouse IgG (ms-IgG) or CDH1 antibody or for 10 d with normal mouse IgG or CDH2 antibody. Each bar represents a mean ± sem of at least three ovaries. Note a marked increase in the percentage of primordial follicles and their development into the primary stage when E15 ovaries were cultured for one additional day. Bars with same letter are not significantly (P > 0.05) different from each other.

CDH2 and CDH1 differentially affect primordial follicle formation in the ovary

We used a neutralization antibody directed against the His-Ala-Val-Asp-Ile and Ile-Asn-Pro-Ile-Ser-Gly-Gln sequences in the extracellular domain 1 of CDH2 and CDH1, respectively (30,31,32,33) to block the CDH1 or CDH2 action during the development of fetal ovaries in vitro. Based on the expression pattern of CDH1 or CDH2 in the postnatal ovaries with respect to primordial follicle formation, E15 ovaries were cultured for 9 or 10 d to ensure the onset of follicle formation. In vitro exposure of cultured ovaries to IgG-Alexa 488 indicated that the IgG reached the center of the ovarian tissue within 16–24 h (data not shown). Consistent with our previous results (19), E15 ovaries cultured in the absence or presence of nonimmune mouse IgG for 9 d had around 10% oocytes transformed into primordial follicles, but primary follicles were not present (Fig. 8C). On the other hand, the percentage of primordial follicles increased significantly in ovaries exposed to the CDH1 antibody (Fig. 8C). After 10 d of culture, the percentage of primordial follicles in ovaries cultured in the absence or presence of nonimmune mouse IgG was approximately 23% (Fig. 8D). Because of the 10-d culture, approximately 4% of primordial follicles also developed into primary follicles (Fig. 8D). The CDH2 antibody showed a modest effect on primordial follicle formation on 9th day of culture (data not shown); however, by 10th day of culture, it markedly reduced the percentage of primordial and primary follicles (Fig. 8D), thus establishing the window of CDH2 action during the initial phase of follicle formation. The antibody-treated ovaries did not show significant chromatopyknosis (an index of cell death), indicating that generalized cytotoxicitiy was not the cause of failure in primordial follicle formation.

Discussion

The present study indicates that Cdh1, Cdh2, CDH1, and CDH2 are spatiotemporally expressed in perinatal hamster ovaries, they exhibit a unique cell-type specific expression with respect to the formation of primordial follicles. Further, this is the first direct evidence to show that CDH1 and CDH2 differentially affect the formation of primordial follicles, and their ovarian expression is differentially affected by FSH during postnatal ovary development. The expression levels of the adhesion molecules significantly affect the assembly of the adhesion junctions at the cell surface and therefore is very important for cell-cell communication and morphogenesis. The transient decrease in CDH1 on P8 against a steady higher level of CDH2 corresponds to the oocyte-somatic cells assembly for the morphogenesis of primordial follicles. Whereas CDH2 may be involved in somatic cell assembly with the oocyte during the formation of primordial follicles, and the apposition of granulosa cells as follicle grow, CDH1 seems to maintain the egg nest for a controlled release of the oocytes for primordial follicle formation. Decreased CDH2 expression and increased CDH1 expression in conjunction with the block in primordial follicle formation in FSH antiserum-treated animals suggest that one of the mechanisms whereby FSH promotes oocyte and somatic cell interaction is by modulating cadherin expression. The steady-state levels of CDH2 relative to TUBB from P8 onwards confirm the granulosa cell only expression of the adhesion protein.

The decrease in CDH1 expression from birth to P7 coincides with the breakdown of the egg nest in preparation of the formation of primordial follicles on P8. Mackay et al. (6) have shown that in the mouse gonad, strong CDH1 staining is present in germ cells and somatic cells in contact with germ cells. CDH1 immunoreactivity in germ cells declines by E16 when ovigerous cords dissociate as a prelude to follicle formation. CDH1 staining reappears transiently on E17 before waning on E19. In the developing pig ovary, CDH1 expression is very high in fetal and neonatal ovaries and declines markedly with maturity (7). Taken together, it seems that a decline in CDH1 expression is necessary for the formation of primordial follicles in the hamster as well.

The differential regulation of CDH2 and CDH1 expression by FSH suggests that cadherin expression in postnatal ovarian cells is controlled by FSH and confirms our previous findings that perinatal hamster ovaries respond to FSH stimulation (17,19). Machell and Farookhi (43) have shown that 48-h eCG treatment of prepubertal rat ovaries results in an increase in Cdh1, but not Cdh2 expression. However, Cdh2 expression increases significantly 24 h after human CG treatment of eCG-treated rats. In contrast, Sundfeldt et al. (44) have reported an abrupt decrease in ovarian Cdh1 protein 4 h after human CG administration to eCG-treated immature rats. In cultured mouse Sertoli cells, FSH is capable of increasing Cdh2 mRNA levels by 2-fold (45). Further, both FSH and estradiol are required to achieve a maximal Cdh2 mRNA levels in cultured Sertoli cells (45). FSH and estradiol have been shown to stimulate Cdh2 mRNA levels in rat granulosa cells in vitro (3,10). MacCalman et al. (9,46) reported that estradiol caused a rapid and significant increases in Cdh1 and Cdh2 mRNA levels in immature mouse ovaries. All these lines of evidence suggest that ovarian expression of Cdh1 and Cdh2 is affected by folliculotropic hormones. Estradiol stimulates the formation and development of primordial follicles in the hamster (19).

The results of the antibody inactivation of CDH1 and CDH2 clearly indicate that these cadherins play a critical differential role during the formation and development of primordial follicles. Antibody neutralization of CDH1 in the explants of sex-indifferent mouse gonads in vitro results in a significant reduction in germ cell number and altered location of germ cells in the gonads (47). The transient decline in CDH1 expression shortly before the formation of primordial follicles in the hamster ovary corroborates similar findings reported for the mouse (6) and human fetal ovary (8) and leads to the speculation that the reduction in CDH1 expression or function in the oocytes is essential for the breakdown of the egg nests so that individual oocytes can assemble with adjacent pregranulosa cells and migrate as primordial follicles. The increase in CDH1 expression (present study) with a corresponding decline in primordial follicle formation (17) in FSH antiserum-treated P8 hamsters, and an increase in the percentage of primordial follicles in CDH1 antibody-treated ovaries in vitro, further support this contention.

The unique expression of CDH2 in postnatal hamster ovaries suggests that CDH2 gene expression shifts from the oocytes to adjacent granulosa cells. It seems that at the initial stage of follicle formation, oocytes make the apposing somatic cells immobile with the help of CDH2. This link may help commit the somatic cells into the granulosa cell lineage, which then adhere to each other, deposit the BM, and become primordial follicles. It is tempting to speculate that oocytes may trigger the expression of CDH2 in pregranulosa cells for potential anchoring by secreting some factor(s), which can affect somatic cells in the immediate vicinity. This speculation stems from the fact that somatic cells further away from the oocyte clusters do not express CDH2. The disappearance of CDH2 expression in the oocytes with the formation of follicular structure suggests that once committed cells are encased in the BM and are adhered to each other with the adhesion molecule, the necessity for the oocytes to produce CDH2 ceases. The results of the present study also suggest that the BM is deposited as soon as undifferentiated somatic cells commit to form pregranulosa cells, and these cells must be encased by the BM to become follicular granulosa cells. However, CDH1 in the oocytes may serve as the adhesion molecule to anchor the granulosa cell processes to the oocyte membrane. With the growth of follicles, the oocyte-granulosa cell contact is stabilized further with the deposition of the zona proteins. The presence of CDH2 in granulosa cells has been shown in many species (10,13,14,46). CDH2 is expressed predominantly in the germ cells of human fetal ovaries in the first trimester, but in the newly formed primordial follicles CDH2 expression, is confined to the pregranulosa cells surrounding the oocyte (8). The cords of somatic cells between germ cell clusters show no CDH2 expression (8). CDH2 plays an important role in establishing cell-to-cell communication by facilitating gap junction formation in many tissues (48,49,50). Gap junctions are essential for maintaining cellular communications between cumulus cells and the oocytes and between granulosa cells in follicles (12,18,51,52,53,54,55). Peluso (56) has shown that CDH2 homophilic binding in granulosa cells results in activation of fibroblast growth factor-receptor and prevention of a sustained release of intracellular Ca2+, which triggers granulosa cell apoptosis. Members of the TGFB family, which functionally interact with CDH2 (57), are also expressed in developing ovaries and regulate the somatic-germ cell interaction (21,29,58,59). The preferential interaction of CDH1 or CDH2 with CTNNB1 in hamster ovarian cells suggests that Wnt/Frizzled signaling pathways may also be involved in the formation and development of primordial follicles. CTNNB1 is the main component in many development-associated signaling pathways (16).

In summary, the results of the present study suggest that differential cell-type specific expression of CDH1 and CDH2 is functionally related to somatic cell assembly with the oocytes to form primordial follicles. Further, FSH regulation of primordial follicle formation may be modulated by the action of cadherins. The results also provide evidence for the first time that although CDH1 may control the temporal migration of the oocytes from the egg nests, CDH2 expression and action are important for the adherence of pregranulosa cells to form primordial follicles.

Footnotes

This work was supported by the National Institute of Child Health and Human Development, National Institutes of Health, Grant R01-HD38468 and by a grant from the Olson Center for Women’s Health Foundation at University of Nebraska Medical Center (S.K.R.).

Disclosure Summary: The authors have nothing to disclose.

First Published Online March 10, 2010

Abbreviations: BM, Basement membrane; CDH1, E-cadherin; CDH2, N-cadherin; CTNNB1, β-catenin; E, embryonic day; eCG, equine chorionic gonadotropin; JUP, γ-catenin; MVH, mouse vasa homolog; NCBI, National Center for Biotechnology Information; P, postnatal day; q, quantitative; TUBB, β-tubulin; WGA, wheat germ agglutinin.

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