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research-article |
Department of Biochemistry and Molecular and Cellular Biology,3 Vincent T. Lombardi Comprehensive Cancer Center,4 Georgetown University Medical Center, Washington, DC 20057
ABSTRACT
Intraovarian growth factors play a significant role in the regulation of follicular selection and growth. In this study, the presence and localization of all members of the family of platelet-derived growth factors (PDGF) and receptors (PDGFR) were identified and characterized in the rat ovary. In addition, a role was identified for members of this family in contributing towards growth of preantral follicles. Real-time PCR revealed the presence of mRNA for all platelet-derived growth factors (Pdgfa, Pdgfb, Pdgfc and Pdgfd) and receptors (Pdgfra and Pdgfrb) in the rat ovary from birth until 4 wk. In situ hybridization and immunohistochemistry were utilized to identify cell-type expression of PDGFs and PDGFRs in rat ovaries from birth until 4 wk. Shortly after birth, expression of PDGFRA and PDGFC was observed in and around oocyte clusters, and PDGFRB in stromal cells surrounding oocyte clusters. All members were identified in oocytes of primordial and primary follicles, and in cells of the theca layer of primordial to antral follicles. PDGFRA and PDGFA were also localized to some granulosa cells of secondary and antral follicles in ovaries from rats at Days 20 and 24. Thus, localization data suggest both theca-theca and theca-granulosa cell interactions of PDGFs and receptors. Preantral follicles cultured in vitro over 5 days in serum-free medium plus recombinant PDGFAA, PDGFAB, or PDGFBB increased in follicle diameter by 18.32% ± 2.18%, 17.72% ± 2.3%, and 17.6% ± 1.81%, respectively, representing significantly greater increases than for follicles incubated in serum-free medium alone (11% ± 1.57%), and suggesting a role for these growth factors in positively influencing early follicle growth.
follicle, granulosa cells, growth factors, ovary, theca cells
Ovarian follicular development is a complicated process involving endocrine, paracrine, and autocrine signals. Folliculogenesis commences within a few days after birth in the rodent when somatic cell proliferation and migration leads to the enclosure of individual oocytes by a single layer of squamous pregranulosa cells and an intact basal lamina [1, 2], thus forming the primordial follicle. Follicle cells and oocyte then become quiescent until their recruitment into the pool of growing follicles and subsequent progression through preantral (primary and secondary follicles) and antral stages [3, 4]. Initiation of follicle growth to the primary stage is observed by an increase in oocyte size and transformation of granulosa cells from squamous to cuboidal. A layer of theca cells becomes observable in the late primary or early secondary stage [5, 6]. During the secondary stage, granulosa cells proliferate to form two to seven layers around the oocyte, and a well-defined thecal layer is formed. At this stage of follicle development, FSH receptors are expressed by granulosa cells and LH receptors are expressed by theca interna cells [7–9]. Antral follicles become apparent when the follicle diameter reaches about 200–250 µm (rodent) [7, 10], at which stage fluid-filled patchy spaces become apparent within the granulosa cell layers. These eventually coalesce to form a crescent-shaped cavity, the antrum [7, 10].
While FSH is essential for growth beyond the antral stage, intraovarian factors are responsible for the early stages of follicle growth. The early signals that initiate follicle growth are still poorly understood but may depend on changes in both inhibitory and permissive factors within the microenvironment of an individual follicle and on signals originating from the oocyte [4, 7]. It is clear that continued early growth of the follicle requires bidirectional communication between granulosa cells and oocytes as well as granulosa and theca cells [7, 11–13]. Because these early follicles might potentially be utilized to increase reproduction efficiency in humans, domestic animals, and endangered species, it is important that the factors involved in the early stages of follicle growth are identified. Two members of the transforming growth factor-ß (TGFB) superfamily, growth differentiation factor-9 (GDF9) and bone morphogenetic protein 15 (BMP15), are produced by oocytes of follicles at all stages of development from primary through ovulation [14, 15] and are essential for follicle growth beyond the primary stage [16–20]. Kit ligand (also known as steel factor or stem cell factor) is synthesized by granulosa cells [21, 22] and binds to receptors on the oocyte, leading to growth of the oocyte, and is essential for the primordial to preantral follicle development [23–27]. Kit ligand also binds to receptors on theca cells and promotes the formation and proliferation of theca cells [28]. Similarly, keratinocyte growth factor (KGF) (also called fibroblast growth factor-7) promotes the survival, growth, and differentiation of medium preantral follicles in vitro [29].
Platelet-derived growth factor (PDGF) has been demonstrated to stimulate in vitro proliferation of theca cells from antral follicles from both rat [30] and pig [31–33] while inhibiting thecal LH-induced steroid hormone synthesis [32]. Transcripts of the 4 Pdgf isoforms (Pdgfa, Pdgfb, Pdgfc, and Pdgfd) have been identified in the human ovary [34], but as yet there are insufficient data to indicate whether this is also true for the rodent ovary. The biological effects of the platelet-derived growth factors are mediated via two structurally similar tyrosine kinase receptors, PDGF receptor alpha (PDGFRA) and PDGF receptor beta (PDGFRB) [35, 36]. PDGFA, PDGFB, and PDGFC bind to PDGFRA, while PDGFB and PDGFD bind to PDGFRB [34, 37]. PDGF is a dimeric molecule and consists of either homodimers (AA, BB, CC, DD) or a heterodimer (AB) that bind two receptors simultaneously, resulting in receptor activation and subsequently inducing a wide variety of cellular responses, including proliferation, survival, and chemotaxis [36]. PDGFs and PDGF receptors are often expressed by neighboring cell types, suggesting a paracrine interaction.
Ovarian expression and localization data for PDGFs and PDGFRs are limited. PDGFRB has been localized to the theca cell layer of adult porcine ovaries [32] and PDGFRA to theca cells and weakly to basal granulosa cells of porcine antral follicles [38]. PDGFA has been observed in some granulosa cells of porcine follicles [38] and in immortalized mouse granulosa cells [39], as well as in the basal lamina of the avian ovarian follicle [40]. In human females undergoing IVF procedures, PDGFs have been identified in follicular fluid [41–43] and PDGFA observed in granulosa cells from large antral follicles [44]. However, a thorough study of the presence, localization, or ontogeny of all the PDGFs and PDGFRs has not been undertaken. In view of the properties of PDGF in promoting cell proliferation, survival, and chemotaxis, it is of interest to determine whether this family of growth factors contributes toward follicle development. Thus the aims of this study were to identify and characterize the presence and localization of all platelet-derived growth factors and receptors in the ovary and to explore a possible role for members of this family in the development of preantral follicles.
Sprague-Dawley rats were maintained in the Animal Care Facility at Georgetown University on a 12L:12D cycle. The Institutional Animal Care and Use Committee of Georgetown University approved all procedures. For ontogeny studies of levels of Pdgf and Pdgfr mRNA in ovaries, several rats at each age group, from newborn to Day 28, were killed at the same time of day (morning) and ovaries for each age group combined for processing. For studies on effects of gonadotropin on Pdgf and Pdgfr mRNA, Day 26 immature rats were injected with 5 I.U. equine chorionic gonadotropin (eCG) and four rats killed at specified time points after injection. In these studies, one ovary from each animal was processed for real-time PCR, and the other ovary placed in 10% neutral buffered formalin (10% formaldehyde solution; EMD Chemicals, Inc., Gibbstown, NJ) for 24 h and processed for immunohistochemistry. Replicate ovaries at each time point were processed separately for real-time PCR.
Preparation of RNA and reverse transcription. After dissection ovaries were removed and placed in RNAlater (Ambion Inc., Austin, TX), while surrounding fat and ovarian bursa were removed. Ovaries were frozen in dry ice and stored at –80°C. Ovarian RNA was prepared using the QIAGEN RNeasy Mini Kit (QIAGEN, Inc., Valencia, CA) and RNA stored at –80°C. Ovarian RNA was reverse transcribed to cDNA using the Taqman Reverse Transcription reagents (Applied Biosystems, Foster City, CA) and random hexamers. The reverse transciption reaction was performed in a BioRad iCycler (BioRad, Hercules, CA) at 25°C 10 min, 37°C 30 min, and 95°C 5 min. Reaction products were stored at –20°C.
Real-time PCR. For real-time PCR, 4 µl of equal dilutions of each reverse transcription reaction product was added to 5 µl 2X SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA) and made up to a final volume of 10 µl with primers and water. The concentration of each set of primers was determined during prior validation experiments using a series of dilutions of each primer set. Forward and reverse 18S rRNA and Pdgfb primers were used at 200 nM, forward and reverse Pdgfa, Pdgfc, Pdgfd, and Pdgfra primers were used at 300 nM, and forward and reverse Pdgfrb primers were used at 600 nM final concentrations. Primers used were: Pdgfa forward 5'- CCC CGG GAG TTG ATC GA -3', reverse 5'- GGT TTG TCT CCA AGG CAT CCT -3'; Pdgfb forward 5'- CTC CAT CCG CTC CTT TGA TG -3', reverse 5'- CAG AAT GTG CTC GGG TCA TGT -3'; Pdgfc forward 5'- CTC GGG CTG AGT CCA ACC T -3', reverse 5'- TGC ACT CCG TTC TGC TCC TT- 3'; Pdgfd forward 5'- CAA CAG CTA CCC GAG GAA CCT -3', reverse 5'- TGG TCA AAG GCC AGC TGT ATC -3'; Pdgfra forward 5'- TTC CCC TGC CAG ACA TTG AC -3', reverse 5'- ATG GCG CTC TCT TCC GAA GT -3'; Pdgfrb forward 5'- GTG AGC GGA AGC GCA TCT A -3', reverse 5'- TCA CTC GGC ATG GAA TCG T -3'; 18S rRNA forward 5'- AGG ATC CAT TGG AGG GCA AG -3', reverse 5'- AAC TGC AGC AAC TTT AAT ATA CGC TAT T -3'. Reaction conditions were 50°C for 2 min, 95°C for 10 min, and 40 cycles of 95°C for 15 sec and 60°C for 1 min using an ABI PRISM 7700 Sequence Detection System (Applied Biosystems, Foster City, CA). CT values (the PCR cycle number at which the PCR fluorescence product is detectable above an arbitrary threshold) for each reaction were determined using ABI PRISM SDS 2.1 analysis software. Quantification of results was determined using the delta delta CT method [45]. Briefly, Pdgf and Pdgfr mRNA levels were normalized to 18S rRNA levels for each quadruplicate sample to compensate for errors in total RNA amounts (Delta Ct). Fold differences in amounts of mRNA per µg total ovarian RNA were determined by comparison of each Delta Ct with an arbitrary control (Day 0 or eCG time–0) (Delta Delta Ct), and the relative value (RQ) determined by raising 2 to the power of the negative Delta Delta Ct value [45]. Single and specific PCR products, as well as lack of contamination or primer dimer formation, were determined by a heat dissociation curve assay at the end of the 40 cycles. Using a 386-well plate, each real-time PCR reaction included quadruplicate samples of mRNA from ovaries from rats at each age, or from one ovary from each of three rats at each time point after eCG injection; thus, one real-time PCR reaction yielded complete data for levels of mRNA for each of the 4 Pdgfs, 2 Pdgfrs, and 18S rRNA. To determine equal efficiency of reactions for Pdgf or Pdgfr and 18S rRNA, each reaction also included a standard curve of log of transcript concentrations versus CT, based upon a number of dilutions of ovarian cDNA prepared for this purpose; each of the seven real-time PCR reactions yielded efficiencies of 95%–100%.
Statistics. For studies of levels of mRNA in response to eCG treatment, comparisons of the x-fold change in Pdgf isoforms and receptors at each time point compared with eCG time–0 were performed using specific contrasts within a generalized estimating equations (GEE) analysis [46]. This analysis calculated the appropriate variance, taking into account the correlation between replicates from the same ovary. A Bonferroni adjustment [47] for significance was utilized such that any P values between 0.05 and 0.0029 were considered interesting, and P < 0.0029 was considered significant. Additionally, a 2-sided binomial test of whether the proportion of time points with fold-changes greater than eCG time–0 was performed for each isoform and receptor. If the P < 0.05, then the isoform or receptor was considered to significantly change following treatment.
Reverse transcription and preparation of riboprobe template cDNA. Ovarian RNA was reverse transcribed to cDNA using the SUPERSCRIPT First-Strand Synthesis System for RT-PCR (Invitrogen Corporation, Carlsbad, CA). Reaction products were stored at –20°C. Complementary DNA templates for the generation of antisense Pdgfb, Pdgfc, and Pdgfd riboprobes were prepared by a PCR reaction using primers specific for each growth factor with the addition of the primer sequence for the T7 promoter to the reverse primer. Primers used were: Pdgfb forward 5'- AAT CGC TGC TGG GCG CTC TT -3', reverse 5'- CCA AGC CTT CTA ATA CGA CTC ACT ATA GGG AGA CTA GGC TCC GAG GGT C -3'; Pdgfc forward 5'- ACA AGG AAC AGA ACG GAG TGC A -3', reverse 5'- ATC GTA ATA CGA CTC ACT ATA GGG TGC CAT CGA TCT GGC TCT AGG -3'; Pdgfd forward 5'- AGG AAG ATG GTG TGG CCA TAA GG -3', reverse 5'- ATC GTA ATA CGA CTC ACT ATA GGG CAT CAT CAT TGA GCC TGT CCA GG -3'. For each reaction, 2 µl of ovarian cDNA were incubated with 1X reaction buffer (Invitrogen Life Technologies, Carlsbad, CA), 2 mM MgSO4 (Invitrogen Life Technologies, Carlsbad, CA), 0.4 mM each of dATP, dTTP, dCTP, and dGTP (Invitrogen Life Technologies, Carlsbad, CA), 2 units PLATINUM Taq High Fidelity (Invitrogen Life Technologies, Carlsbad, CA) and molecular grade water to a final volume of 50 µl. The PCR reaction was performed in a BioRad iCycler at 94°C 2 min and 30 cycles of 94°C for 30 sec, 56°C for 30 sec, and 72°C for 1 min. Amplification products were stored at –20°C. cDNA was purified from primers and reaction components using a Microcon 100 column and eluted in 50 µl molecular grade water. Concentration was determined by spectrophotometry at 260 nm.
Preparation of riboprobes. Riboprobes were prepared using the DIG RNA Labeling Kit (Roche Diagnostics GmbH, Mannheim, Germany). The RNA probes were purified to remove unincorporated nucleotides using a Chroma Spin-100 column (BD Biosciences, Palo Alto, CA) washed with DEPC-water. Briefly, 17 µl RNase-free STE buffer (500 mM NaCl, 1 mM EDTA, 20 mM Tris-HCl, pH 7.5) and 20 µg glycogen (Roche Diagnostics GmbH, Mannheim, Germany) as carrier to maximize yield were added to the riboprobe preparation and the total volume (40 µl) run through the Chroma Spin-100 column, yielding 40 µl of RNase-free probe in DEPC-water. Concentration of probes was ascertained by spectrophotometry at 260 nm. RNase inhibitor (Roche Diagnostics GmbH, Mannheim, Germany) was added at 1 U/µl final volume and probes aliquoted and stored at –80°C. The integrity of the probes was determined by electrophoresis on a 1% formaldehyde denaturing agarose gel followed by Dot blotting.
In situ hybridization. A number of methods were followed and positive signal identified with either NBT/BCIP or DAB chromogen. Paraformaldehyde-fixed ovaries were washed by 2 x 30 min washes in phosphate buffered saline (PBS) and dehydrated through a series of incubations in increasing concentrations of ethanol, followed by clearing with xylene. Ovaries were then embedded in paraffin and stored at 4°C. Ovaries were sectioned (7 µm), cleared of paraffin in xylene, and rehydrated through decreasing concentrations of ethanol (100% and 50%). After washing in DEPC-treated water, sections were incubated in PBS at 37°C for 5 min, followed by 10 min in 10 µg/ml Proteinase K (Sigma Chemical Co, St. Louis, MO) in PBS at 37°C. Sections were washed in DEPC-treated water and then in 0.2N HCl in DEPC-water at room temperature for 15 min. Tissue was acetylated by submersing for 15 min in 0.25% acetic anhydride (Sigma Chemical Co., St. Louis, MO) in 0.1 M triethanolamine (Sigma Chemical Co., St. Louis, MO) pH 8.0 at room temperature. Tissue RNA was denatured by incubating slides 15 min in 2X SSC at 70°C. Riboprobes were denatured at 70°C for 5 min and 10 ng probe added to 100 µl hybridization solution containing 50% formamide (Electron Microscopy Sciences, Ft. Washington, PA), 10% dextran sulfate (Intergen, Purchase, NY), 4X SSC, 1X Denhardt's Solution (Quality Biological Inc., Gaithersburg, MD), 2.5 mg/ml yeast tRNA (Roche Diagnostics GmbH, Mannheim, Germany), and 0.5 mg/ml heat denatured herring sperm DNA (Roche Diagnostics GmbH, Mannheim, Germany). Hybridization occurred overnight at 50°C. The following day, slides were rinsed in 2X SSC and washed twice in 2X SSC, followed by two washes each at 55°C in 1X SSC, 0.5X SSC, and 0.1X SSC. Sections were blocked in 2% heat inactivated sheep serum (Sigma Chemical Co., St. Louis, MO) in TN Buffer for 30 min at room temperature. Slides were then incubated at 4°C overnight in 1:500 dilution of anti-Digoxigenin-AP Fab fragments (Roche Diagnostics GmbH, Mannheim, Germany) in TN Buffer with 1% heat inactivated sheep serum. Slides were rinsed and washed in 2X SSC, followed by 2 washes in TN Buffer, and 2 washes in TNM Buffer at room temperature. Hybridization was visualized after incubation in 45 µl NBT (Roche Diagnostics GmbH, Mannheim, Germany), 35 µl BCIP (Roche Diagnostics GmbH, Mannheim, Germany) and 1 mM levamisole (Sigma Chemical Co., St. Louis, MO) in TNM buffer in the dark. The color reaction was stopped by placing slides in Stop Buffer (10 mM Tris-HCl, pH8, 1mM EDTA). Slides were washed in water, allowed to air dry, mounted with GVA mounting solution (Zymed, S. San Francisco, CA), and cover slipped.
Alternatively, positive signal was visualized after incubating sections with a biotin-labeled anti-digoxin antibody (InnoGenex, San Ramon, CA) followed by incubation in peroxidase conjugated streptavidin (BioGenex, San Ramon, CA) and DAB chromagen (Zymed, S. San Francisco, CA). Sections were counterstained with hemotoxylin (Fisher Scientific, Fair Lawn, NJ).
Rats were killed and the ovaries removed and either immersed in Tissue-Tek O.C.T. Compound (Sakura Finetek USA Inc., Torrance, CA) and snap frozen in liquid nitrogen followed by storage at –80°C, or placed in 10% neutral buffered formalin for 24 h and processed for paraffin embedding. Frozen sections (5 µm) were fixed in ethanol for 15 min, rinsed in water, and immersed in PBS pH 7.6 for 15 min. Endogenous peroxidase activity was quenched in 0.3% H2O2 (Fisher Scientific, Fair Lawn, NJ) in PBS for 5 min at room temperature and then washed in PBS. Sections were blocked for 45 min at room temperature with 10% normal goat serum (Vector Laboratories, Burlingame, CA) in PBS, washed, and incubated with antibody specific for PDGFRB (sc432; Santa Cruz Biotechnology, Santa Cruz, CA) diluted 1:200 in 1% BSA in PBS overnight at 4°C. Sections were then washed with PBS, incubated with biotinylated anti-rabbit secondary antibody (Vector Laboratories Inc., Burlingame, CA) diluted 1:200 in 1% BSA in PBS for 1 h at room temperature and washed with PBS. Immunoreactivity was visualized after 20 min incubation in peroxidase conjugated streptavidin at room temperature followed by peroxidase reduction of AEC substrate (Zymed, San Francisco, CA). Sections were counterstained with hemotoxylin.
For paraffin-embedded ovaries, 3 µm sections were deparaffinized in xylene for 2 x 5 min, followed by rehydration in decreasing concentrations of ethanol (100%, 90%, 70%, and 50%) and finally in water for 5 m. Antigen retrieval was performed in Retrievit (Biogenex, San Ramon, CA) at pH 5.5 (PDGFRA, PDGFAA, PDGFC) or pH 6 (PDGFRB, PDGFB) in a microwave at 20% maximum for 10 min. After cooling at room temperature for 30 min, sections were washed in PBS and endogenous peroxidase activity was quenched in 0.3% H2O2 in PBS for 5 min at room temperature. Sections were washed in PBS and blocked for 45 min at room temperature with 10% normal goat or rabbit serum (Vector Laboratories, Burlingame, CA) in PBS, washed and incubated in a 1:100 dilution of antibody specific for PDGFRA (sc338; Santa Cruz Biotechnology, Santa Cruz, CA), PDGFRB (sc432; Santa Cruz Biotechnology, Santa Cruz, CA), PDGFAA (BioDesign International, Saco, Maine), PDGFB (sc7878; Santa Cruz Biotechnology, Santa Cruz, CA) or PDGFC (sc-18228; Santa Cruz Biotechnology, Santa Cruz, CA) in 1% BSA in PBS at 4°C for 24 to 48 h. Sections were washed with PBS and incubated with biotinylated secondary antibody (Vector Laboratories, Burlingame, CA) diluted 1:200 in 1% BSA in PBS for 1 h at room temperature. Sections were washed with PBS, incubated in peroxidase-conjugated streptavidin for 20 min at room temperature and immunoreactivity visualized by peroxidase reduction of AEC substrate. Sections were counterstained with hemotoxylin. Alternatively, in some experiments, antigen retrieval was omitted and sections incubated with antibody at 4°C for 4 days.
Plate preparation. Prior to use, 96-well cell culture plates were treated with Poly(2-hydroxyethyl methacrylate) (polyHEMA) (Sigma Chemical Co., St. Louis, MO) to prevent attachment of follicle cells to the well surface. 100 µl of a 1:10 preparation of polyHEMA:95% ethanol was dispensed into each well and allowed to evaporate overnight in a tissue culture hood under ultraviolet irradiation.
Follicle isolation. Ovaries were excised from immature Sprague-Dawley rats aged between postnatal Days 11 and 15. One animal was killed per experiment and the ovaries dissected and placed in a 100-mm culture dish with 10 ml pre-warmed alpha-MEM medium containing GlutaMAX, ribonucleosides, and deoxynucleosides (Gibco Invitrogen Corporation, Grand Island, NY) supplemented with 0.3% BSA (Fisher Scientific, Fair Lawn, NJ) and 100 units/ml penicillin and 100 µg/ml streptomycin (Gibco Invitrogen Corporation, Grand Island, NY). Preantral follicles with mean diameters between 120 and 200 µm were mechanically dissected using two 1-ml syringes (Becton Dickinson and Company, Franklin Lakes, NJ) with size 27G 1/2 needles (Becton Dickinson and Company, Franklin Lakes, NJ) and collected in a basal culture medium consisting of alpha-MEM (with GlutaMAX, ribonucleosides, and deoxynucleosides), 0.1% BSA, 50 µg/ml freshly prepared ascorbic acid (Acros Organics, NJ), 1 x ITS (10 ng/ml insulin, 110 ng/ml pyruvate, 5.5 µg/ml transferrin, 0.67 µg/ml selenium; Gibco Invitrogen Corporation, Grand Island, NY), 100 units/ml penicillin. and 100 µg/ml streptomycin. Isolated follicles were incubated in a humidified incubator gassed with 5% CO2 in air at 37°C until sufficient follicles were isolated for culture.
Follicle culture. Follicles were transferred to polyHEMA-treated 96 well culture plates (Corning Incorporated, Corning, NY), each well containing 50 µl basal culture medium as described earlier. After transfer, a further 50 µl of basal culture medium plus treatments were added to each well. The final concentration of treatments were: 100 ng/ml recombinant rat PDGFAA (R&D Systems, Inc., Minneapolis, MN), 100 ng/ml recombinant rat PDGFAB (R&D Systems, Inc., Minneapolis, MN), 100 ng/ml recombinant rat PDGFBB (R&D Systems, Inc., Minneapolis, MN), 15 µM AG1296 (Calbiochem, La Jolla, CA) or 1 I.U./ml recombinant human FSH (National Hormone and Peptide Program, Torrance, CA). Approximately half of the culture medium was removed daily and replaced with freshly prepared medium and treatments. Follicles were incubated in a humidified incubator gassed with 5% CO2 in air at 37°C for 5 days.
Measurements. Follicles were imaged daily using a Nikon Eclipse TE-300 Inverted Microscope System in a 5% CO2 gassed chamber. At the completion of the culture period, serial images of each follicle were viewed and follicle diameters measured using MetaMorph (version 5.0.3) Image Acquisition & Analysis software (Universal Imaging Corp). Measurements consisted of 2-dimensional diameters taken at perpendicular angles, and the mean determined from these. Follicles of 120–200 µm with normal morphological appearance at Day 0, i.e., a central spherical oocyte, with or without a small amount of attached stroma [48, 49] and that remained intact and exhibited growth over the 5-day culture period [50–53] were analyzed. With the exception of the positive control group of follicles cultured in the presence of FSH, a similar percentage of follicles from each group showed growth over 5 days. A higher percentage of follicles cultured in the presence of FSH exhibited growth over the culture periods. Sample sizes for the 5-day culture period were: control n = 7, PDGFAA n = 9, PDGFAB n = 9, PDGFBB n = 6, AG1296 n = 7, and FSH n = 14.
Statistics. Results are expressed as average percentage increase in mean follicle diameter compared with Day 0 ± SEM for each group. Statistical significance between average percentage follicle growth at Day 5 was determined by ANOVA followed by t-test analysis of each group in comparison with average percentage increase in follicle diameters of the control group.
Expression of PDGF Receptors in the Ovary
Messenger RNA for Pdgfra (Fig. 1A) and Pdgfrb (Fig. 1B) was detected in rat ovaries from newborn to Day 28. Both receptors follow a similar pattern of expression, with the amount of mRNA for each receptor per µg total ovarian RNA apparently peaking at Days 4–8 after birth at a 2-fold increase over levels at birth. It must be noted, however, that absolute amounts of mRNA for each of these receptors were not determined from these studies, and thus a comparison between the quantities of each message cannot be made.
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Immunohistochemistry, using antibodies to PDGFRA and PDGFRB, was employed to determine which cells within the ovary express the receptors. Observations are summarized in Table 1. Using paraffin-embedded sections, both with and without antigen retrieval, PDGFRA immunoreactivity was observed in ovaries from Day 4 rats in regions of oocyte clusters, localized to either the oocyte or to cells interspersed between oocytes (Fig. 2, A and B). Similarly, PDGFRA staining was apparent in primordial follicles, localized to either the oocyte or to pregranulosa cells (Fig. 2, A–C), and to the oocyte of primary follicles (Fig. 2C). Positive immunostaining was also observed in the theca layer of secondary and antral follicles (Fig. 2, C, E, F, H, and I). In some of these follicles, it is possible that immunopositive cells are cells of the thecal vasculature, while in other follicles, and particularly in antral follicles of rats at Day 20 and in the absence of antigen retrieval, theca cell localization is clear (Fig. 2F). Particularly in ovaries from rats at Day 20 and older, PDGFRA immunoreactivity was apparent in granulosa cells of some secondary and antral follicles (Fig. 2, D, E, G, and H). In frozen sections and in the absence of antigen retrieval, PDGFRB was observed in cells comprising the cord-like structures that separate clusters of oocytes in ovaries from newborn rats (Fig. 2K). By Day 8 post-birth, the receptor was observed in cells surrounding primordial and primary follicles, and in well-differentiated thecal cells of secondary follicles (Fig. 2M). Strong thecal and interstitial cell immunoreactivity was observed in further stages of follicular growth (Fig. 2, N and O). In paraffin-embedded sections, positive staining was also observed in the oocyte cytoplasm of primordial and primary follicles both in the absence and inclusion of antigen retrieval (Fig. 2L).
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Expression of PDGFs in the Ovary
To determine whether the ovary provides a local source for Pdgfs, real-time PCR was employed and message was detected for all 4 Pdgf isoforms (Fig. 3, A–D). These results show a decrease in ovarian message for Pdgfa and Pdgfb per µg total ovarian RNA from Day 2, reaching a nadir by Day 12, and this was maintained until Day 28. Thus, by Day 12 the levels of ovarian message for Pdgfa and Pdgfb per µg total ovarian RNA were 20% and 60%, respectively, of the level of message per µg total ovarian RNA from Days 0 and 2 (Fig. 3, A and B). Levels of Pdgfc mRNA per µg total ovarian RNA varied from Days 0 to 20, and then increased approximately 3-fold by Day 28 (Fig. 3C). In contrast, Pdgfd mRNA levels per µg total ovarian RNA remained relatively constant with a slight cyclical pattern of increase at Day 8 followed by a decrease in ratio (Fig. 3D). Absolute amounts of mRNA for each of these ligands were not determined from these studies, and thus comparisons between each message cannot be made.
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In order to determine the cellular sources of the ovarian Pdgf messages detected by real-time PCR, in situ hybridization of rat ovaries with riboprobes specific for Pdgfb, Pdgfc, and Pdgfd was performed. Message for Pdgfb (Fig. 4, A and B), Pdgfc (Fig. 4, C and D), and Pdgfd (Fig. 4, E and F) was observed in theca cells of secondary follicles in ovaries collected from 12- and 16-day-old rats. Pdgfd message was also localized to oocytes of primordial and primary follicles (Fig. 4, E and F).
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Several methods of immunohistochemistry were explored in an endeavor to yield clear data to identify cellular expression of the platelet-derived growth factor proteins. Observations are summarized in Table 1. In paraffin-embedded sections of rat ovaries at birth (Day 0) (Fig. 5A) and Day 4 (Fig. 5, B and C), PDGFAA immunoreactivity was observed within clusters of oocytes, either on the oocyte membrane or in cytoplasmic protrusions from surrounding somatic cells. At Day 4, PDGFAA immunostaining was also apparent in either the oocyte or pregranulosa cells of primordial follicles, and in the oocyte of primary follicles (Fig. 5, B and C). Immunoreactivity was observed in the theca layer of primary and secondary follicles (Fig. 5, D, F, and G) and in antral follicles, particularly in the absence of antigen retrieval (Fig. 5G). PDGFAA was also observed in granulosa cells of some preantral and antral follicles in ovaries from rats aged Days 20 and 24, but not at earlier ages (Fig. 5, E and F). PDGFB immunoreactivity was observed in interstitial cells in the newborn rat ovary (Fig. 5I), and in theca cells of secondary and antral follicles (Fig. 5, J and K). PDGFB immunostaining was apparent in what appeared to be the basement membrane between the granulosa and theca cell layers of a number of secondary follicles (Fig. 5J). At Day 4, PDGFC protein was clearly identified in cells of oocyte clusters and in the oocyte of primordial and primary follicles (Fig. 5, L and M). In ovaries from rats at Day 20 and older, prominent PDGFC immunostaining was observed in cells of the theca layer of secondary and antral follicles, particularly in the absence of antigen retrieval (Fig. 5, N and O).
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Effect of PDGF on Growth of Preantral Follicles In Vitro
The presence of PDGFs and their receptors in follicles at all stages of development raises the question of a role for this family of growth factors in follicle development. To explore this, follicle cultures were employed (Fig. 6). Preantral follicles cultured in serum-free medium increased in size by 11.00% ± 1.57% over 5 days. A positive control group of follicles were incubated in the presence of FSH, as many studies have shown the importance of FSH for the promotion of in vitro growth and survival of preantral follicles. The addition of FSH to the culture medium led to a significant increase in size after 5 days, namely 24.94% ± 2.56%. Addition of each of the 3 dimers of recombinant platelet-derived growth factors, AA, AB, or BB, to the basal culture medium improved growth of follicles significantly over 5 days when compared with controls. Follicles incubated in the presence of PDGFAA increased in mean diameter by 18.32% ± 2.18%. PDGFAB added to the culture medium resulted in follicles with a mean increase in diameter of 17.72% ± 2.30% after 5 days culture, and preantral follicles cultured in the presence of PDGFBB exhibited a 17.60% ± 1.81% increase in mean diameter after 5 days. No significant difference in growth rates was observed between follicle groups treated with the three platelet-derived growth factors (Fig. 6).
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It has been demonstrated that PDGF receptors are located on theca cells of developing follicles. Inhibition of the activity of these endogenous PDGF receptors (PDGFRA and PDGFRB) by the tyrphostin AG1296 resulted in a reduced increase in mean follicle diameter over 5 days (8.03% ± 1.15%) (Fig. 6) when compared with control group of follicles (11.00% ± 1.57%). However, this reduction in growth rate was not statistically significant.
Effect of eCG on Ovarian Pdgf mRNA Levels
To investigate whether FSH affects expression of Pdgfs and Pdgfrs, immature rats were treated with eCG, a nonpituitary hormone with FSH and some LH properties, and ovarian RNA extracted from rats killed at various times after treatment. Levels of mRNA per µg total ovarian RNA for both Pdgf receptors (Fig. 7, A and B) and all 4 isoforms of Pdgf (Fig. 7, C–F) did not vary significantly following eCG treatment. However, it is interesting to note that Pdgfa mRNA levels were reduced after eCG injection, particularly by 48 h post-injection, reaching 59% of levels prior to injection with P = 0.03, although this was not considered significant by the statistical tests utilized for analysis of this study.
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Although many growth factors are present in serum and can therefore affect ovarian functions after vascular delivery to the ovary, a growth factor that plays an important role in follicle development must be expressed by cells of the ovary so that its expression and downstream effects can be locally regulated. Thus the presence of mRNA for all Pdgfs and their receptors in the ovary from birth until Day 28 is of interest and suggests a role for this family of growth factors during development of the ovary and of the follicle.
Messenger RNA for Pdgfra, Pdgfrb, Pdgfa, and Pdgfd relative to total ovarian RNA was observed to increase in the first few days after birth. Although the ratio of message to total ovarian RNA decreased after this time, as does the postnatal ratio of Pdgfb message, it is not clear whether the absolute levels of mRNA for the Pdgfs and Pdgfrs actually increased, decreased, or remained constant, due to the ongoing increase in total ovarian RNA during the postnatal growth period of the ovary. Pdgfc is the one exception, as it can be seen from the real-time PCR data that the relative levels of mRNA for this growth factor in comparison with total ovarian RNA increased threefold by 4 wk after birth.
In situ hybridization localized Pdgfb, Pdgfc, and Pdgfd mRNA expression to theca cells and also to the oocyte cytoplasm. Protein expression of PDGFA, PDGFB and PDGFC was observed in theca cells. PDGFB protein also appeared to be present either on theca cells lining the basement membrane or in the basement membrane separating theca cells from granulosa cells. A number of studies have identified platelet-derived growth factors in the extracellular matrix, in the basement membrane at the epithelial/mesenchymal interface of most tissues [54, 55] and as a component of the basal lamina of the avian preovulatory ovarian follicle [40]. PDGFBB has been shown to bind to various extracellular matrix components [56, 57] suggesting that attachment to matrix or cell surface may be alternative fates for these polypeptides.
Protein expression of both receptors was clearly observed in the theca cell layer of follicles and in the oocyte or surrounding cells of early stage follicles. Immunoreactivity of PDGFRA, PDGFAA, and PDGFC was observed in oocyte clusters and in primordial follicles. In oocyte clusters it is not clear whether localization of the proteins is in the oocyte membrane or the areas between oocytes, possibly the cytoplasmic extensions of somatic cells. Likewise, in primordial follicles it is unclear whether the proteins are localized to the oocyte membrane or to pregranulosa cells. In sections of ovaries from rats aged Day 20 and 24, PDGFRA and PDGFAA were also present in granulosa cells of some preantral and antral follicles. It is interesting to note that in the female rat, a dramatic increase in systemic FSH concentration has been observed at anywhere from Days 14 to 16 postpartum [52, 58–60] and this leads to FSH-induced differentiation of granulosa cells [52, 61]. FSH receptors are expressed in granulosa cells of secondary and antral follicles. It is possible that the protein expression of PDGFRA and PDGFAA observed in granulosa cells of rats at Days 20 and 24 occurs in response to FSH signaling to these cells at this stage. However it is not clear why PDGFRA or PDGFAA protein immunostaining is not apparent in all granulosa cells or in all secondary or antral follicles, although the state of the follicle's viability might be a factor. These data are similar in part to recent observations in the porcine ovary, in which PDGFRA and PDGFA protein expression was observed in oocytes of primordial follicles and of antral follicles and in some granulosa cells of antral and mature follicles [38].
In the newborn rat ovary, expression of the PDGFRB protein was localized to stromal cells surrounding clusters of oocytes. At later ages, PDGFRB protein expression was observed in cells surrounding the oocyte of primordial follicles. At the level of magnification allowed by resolution, it could not be determined whether the positive immunoreactivity was localized in the pregranulosa cells or from cells surrounding this layer. However, at subsequent stages of follicle development expression of PDGFRB protein was localized to theca cells. Thus, to be consistent with these observations, it is likely that PDGFRB is expressed by cells surrounding the pregranulosa cells, and raises the possibility that these cells are the precursors to the theca cells that are observable at later stages of follicle development. Although it is generally considered that clearly defined theca cells are not recognizable until the late primary or early secondary stage, or when there are several layers of granulosa cells present, Hirshfield [2, 5] has noted that in the rat, theca cells are evident in very small follicles in the earliest stages of growth and these cells proliferate along with the granulosa cells as the follicle grows. It is possible that it is these cells that are observed in these studies as being immunoreactive for PDGFRB in rat primordial follicles. These expression patterns suggest a possible role for PDGF signaling in the perinatal period and could include proliferation and chemotaxis of stromal cells in the growing ovary.
The observation of immunoreactivity of various platelet-derived growth factors and receptors in the oocyte is interesting. While the immunoreactivity of the oocyte of primordial and primary follicles appears to be quite specific, it is not clear whether the intense oocyte staining observed in follicles at later stages of development are due to the presence of the proteins, or a property of the oocyte in attracting non-specific binding of antibodies. To be sure of such localization, it would be necessary to isolate oocytes and determine by western blotting whether these proteins are really present.
To explore a possible role for PDGFs during follicle development, follicle cultures were employed. For the purposes of this study, to ensure the preservation of the spherical follicle structure during culture so as to allow for normal intrafollicular interactions, it was important to prevent attachment of follicle cells to the plate, and this was done by coating the well surfaces with polyHEMA (2-hydroxyethyl methacrylate), an hydrogel that prevents cellular attachment and spreading [62]. Culture conditions were chosen that would support growth of the follicle but not confound or mask the effects of the growth factors under investigation, especially if the effects of these factors are minimal, as might be expected in a system whereby already numerous factors have been demonstrated to contribute towards early follicle growth, as discussed earlier. This necessitated the elimination of serum and FSH from the culture medium. Ascorbic acid and selenium were added to the medium to maintain follicular stability and protect against free radicals, thereby increasing the chance of follicle survival [49, 63, 64]. Ascorbic acid is known to promote collagen biosynthesis [65] and the addition of ascorbic acid to culture medium results in a significant decrease in the number of follicles undergoing degeneration of the basal lamina [66]. Ascorbic acid and selenium also act as antioxidants [67] and reduce the degree of apoptosis within follicles cultured under serum-free conditions [66]. Thus in the minimal medium chosen for these studies, a percentage of follicles survived and increased in size and it was possible to observe the effects of the exogenously added platelet-derived growth factors, as well as inhibition of the receptors, on follicle growth and survival.
Under the culture conditions used in the current study, preantral follicles grew over 5 days. The percentage increase in size (11.00% ± 1.57%) compares favorably with that observed by McGee et al. [68] who reported a 3.60% increase in diameter of 160–200 µm rat preantral follicles over 2 days in serum-free medium (cf. 5.40% ± 0.87% increase after 2 days in the present study). The addition of FSH to the serum-free medium led to a significant increase in mean follicle diameter after 5 days (24.94% ± 2.56%). Similarly, in a serum-free culture medium supplemented with FSH, Zhao et al. [51–53] observed 14%–20% increases in diameter of 140–160 µm rat preantral follicles after 6 days. Fowler and Spears [69] reported a reduction in size of rat follicles when cultured in serum-free medium, but the addition of FSH to the medium resulted in follicle survival and 16% increase in follicle diameter over 3 days (cf. 13.91% ± 1.56% after 3 days in the current study). Many studies demonstrate the important role of FSH in preantral follicle growth and survival in vitro [70–73].
The addition to the culture medium of a recombinant form of each of three conformations of platelet-derived growth factors, PDGFAA, PDGFAB, or PDGFBB, increased growth of follicles significantly when compared with control conditions. The three dipeptides were used to determine whether either one of the two PDGF receptors, PDGFRA or PDGFRB, was primarily or solely involved in follicle growth. Similar rates of growth were observed with the addition of each of the recombinant proteins, ranging from 17.60% to 18.32% after 5 days, demonstrating that both receptors are active in follicle growth.
Platelet-derived growth factors and receptors are expressed in the ovarian follicle at all stages of development, and this ligand-receptor system is active in inducing follicle growth at the preantral stage, as inhibition of the receptors in vitro by the PDGF receptor inhibitor, tyrphostin AG1296, resulted in a smaller increase in follicle size after 5 days than that achieved by follicles cultured in the presence of control medium alone, although this difference was not statistically significant. It must be noted that although AG1296 potently inhibits signaling of PDGFRA and PDGFRB, at a higher concentration it also inhibits the activity of the related stem cell factor receptor, c-Kit [74].
Thus the data derived from these studies clearly implicates signaling by PDGF and its receptors in contributing toward growth of preantral follicles. This effect could be attributable to a proliferation of the theca cell layer and/or induction of synthesis and/or release of another factor from theca cells, which subsequently influences growth of neighboring granulosa cells. The current studies were not detailed enough to elucidate the mechanism by which PDGF signaling positively affected preantral follicle growth.
Antral follicle growth is dependent on FSH signaling. FSH exerts its effects not only directly but also indirectly through increasing expression of locally expressed growth factors [6]. The present study made a preliminary investigation into the possible interaction of FSH and platelet-derived growth factors and receptors. Although mRNA levels were not significantly regulated by FSH, the subsequent protein levels were not explored. In addition, a possible synergistic effect of PDGF and FSH on follicle growth was not investigated. As expression of PDGFRA and PDGFA has been observed in granulosa cells of rats after, but not before, the age at which systemic FSH levels increase dramatically in the rat, it is possible that the activity of this receptor in granulosa cells influences or is influenced by FSH. Thus it would be of interest to determine the presence of synergism of both FSH and PDGF on in vitro preantral and antral follicle growth in comparison with follicles cultured in the presence of either factor alone. In addition, as these studies utilized follicles from rats aged 11 to 15 days, follicle growth in the presence of the exogenous platelet-derived growth factors might have been limited by activity solely within the theca cell layer, where expression of the receptors has been localized at this age. It would therefore be of interest to observe the effects of recombinant platelet-derived growth factors on in vitro growth of preantral follicles from rats aged 20 days or older to determine whether growth might be further increased by the interaction of growth factors with PDGFRA expressed on granulosa cells at these ages.
In summary, these data demonstrate, for the first time, expression of all PDGFs and PDGF receptors in the ovary of both immature and mature rats. Previous studies have elucidated a role for PDGF in inducing theca cell proliferation in theca cell cultures [30–33]. Thus, platelet-derived growth factors synthesized and released by theca cells may act in an autocrine or paracrine manner on theca cells expressing both receptors. Theca-granulosa cell interactions are also possible in view of granulosa cell expression of PDGFRA and PDGFA. Significantly, a role for PDGFA and PDGFB was identified in contributing towards growth of preantral follicles. A definitive insight into the mechanisms by which these growth factors contribute towards growth of preantral follicles, as well as whether this family of growth factors might interact with FSH in influencing follicle development, remains to be determined.
ACKNOWLEDGMENTS
The authors would like to thank Rebecca Slack, M.S., for statistical analyses, and Robert Koos, Ph.D., and Jodi Flaws, Ph.D., for helpful discussions.
FOOTNOTES
1Supported by NIH grants HD36013 and HD39523. 
Correspondence: 2Leanne Sleer, Georgetown University Medical Center, 3970 Reservoir Rd., Washington, DC 20057. FAX: 202 687 8434; e-mail: sleerl{at}georgetown.edu
Received: 22 August 2005.
First decision: 20 September 2005.
Accepted: 3 November 2006.
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