Biol Reprod Keystone Symposia Conference on Frontiers in Reproductive Biology & Regulation of Fertility.
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BOR - Papers in Press, published online ahead of print March 28, 2007.
Biol Reprod 2007, 10.1095/biolreprod.106.053629
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BIOLOGY OF REPRODUCTION 77, 226–236 (2007)
DOI: 10.1095/biolreprod.106.053629
© 2007 by the Society for the Study of Reproduction, Inc.

The Hedgehog Signaling Pathway in the Mouse Ovary1

Matthew C. Russell, Robert G. Cowan, Rebecca M. Harman, Ashleigh L. Walker, and Susan M. Quirk 2

Department of Animal Science, Cornell University, Ithaca, New York 14853

ABSTRACT

The hedgehog (HH) signaling pathway plays an essential role in the Drosophila ovary, regulating cell proliferation and differentiation, but a role in the mammalian ovary has not been defined. Expression of components of the HH pathway in the mouse ovary and effects of altering HH signaling in vitro were determined. RT-PCR analyses show developmentally regulated expression of sonic (Shh), indian (Ihh) and desert (Dhh) HH in the ovary. Expression is detected in whole ovary, granulosa cells, and corpora lutea. The mRNAs for the two receptors, patched homolog 1 and 2 (Ptch1, Ptch2), and the signal transducer, smoothened (Smo), are also expressed. Immunohistochemistry using an antibody that detects all three HH ligands demonstrated HH protein primarily in granulosa cells of follicles from primary to antral stages of development. Follicles also stained for PTCH1 and SMO in both granulosa and theca cells. Treatment of cultured preantral follicles and granulosa cells with recombinant SHH increased growth and proliferation while treatment with the HH pathway inhibitor, cyclopamine, had no effect. Therefore, activation of HH signaling can increase cell proliferation and follicle growth but is not essential for these processes in vitro. Treatment of granulosa cells with SHH increased levels of mRNA for Gli1, a transcriptional target of HH signaling, while cyclopamine decreased expression. SHH had no effect on production of progesterone by cultured granulosa cells, while cyclopamine increased progesterone production. The results demonstrate a functional HH pathway in the follicle and identify granulosa cells as at least one of the potential targets of HH signaling.

follicle,, follicular development,, granulosa cells,, ovary,, theca cells

INTRODUCTION

Ovarian follicle development is a dynamic process of tissue remodeling involving signaling pathways that are essential for development and patterning in many tissues. In the vertebrate embryo and in Drosophila, the hedgehog (HH) signaling pathway directs the patterning of diverse tissues by regulating cell proliferation and survival, cell fate determination, differentiation, and polarity [1]. HH signaling also controls the maintenance and turnover of adult tissues, including the mammary gland, uterus, hair follicle, gastrointestinal tract, pancreas, and others [1, 2]. In addition, deregulation of the pathway has been implicated in the etiology of several cancers [3]. Despite extensive evidence for the importance of HH signaling in the Drosophila ovary, information about HH signaling in the mammalian ovary is just beginning to emerge [4].

In the ovary of adult Drosophila, HH stimulates the proliferation of somatic stem cells, which generate the population of follicle cells that surround developing gametes [5–7]. HH also directs some of the somatic cells to become polar cells or stalk cells of the egg chamber [5, 6]. In addition to effects on somatic cells, HH signaling promotes the proliferation of germ cells by interaction with other critical pathways [8]. In the Drosophila embryo, HH signaling is necessary for migration of primordial germ cells to the developing gonads [9]. During pupal stages of development, HH signaling modulates interactions between somatic and germ cells necessary for the formation of the adult ovary [10].

While a single HH protein exists in Drosophila, three HH proteins have been identified in mammals; sonic (SHH), indian (IHH), and desert (DHH) [11]. HH ligands are secreted proteins that act as morphogens, signaling to nearby cells in a target field according to a concentration gradient [11]. In mammals, all three HH ligands signal by binding to one of two homologous transmembrane receptors, patched homolog 1 and 2 (PTCH1 and 2), which are thought to function similarly. When PTCH is not bound by HH ligand, it blocks the activity of the transmembrane signal transducer protein, smoothened (SMO). Binding of HH to PTCH removes inhibition of SMO, and downstream signaling occurs [11]. Signaling includes activation of a group of transcription factors, GLI-Kruppel family members 1, 2, and 3 (GLI1, GLI2, and GLI3), which may act as either transcriptional activators or repressors [12]. Transcription of target genes in response to HH signaling includes increased expression of Ptch1 and Gli1. Thus, HH signaling alters expression of genes encoding proteins within the HH pathway, thereby modulating its activity [13–15]. In fact, heterozygous deletion of Ptch1 in mice and inactivating mutations of Ptch in Drosophila result in overactivation of HH signaling [7, 16]. In addition, HH signaling is modulated by HH-induced transcription of the gene for HH interacting protein (Hhip), a secreted protein that binds to HH ligands and prevents their interaction with PTCH [17, 18].

The objective of experiments reported here was to examine expression of components of the HH pathway in the mouse ovary and to test whether the pathway is functional by studying effects of stimulating or inhibiting the pathway on cultured follicles and granulosa cells. End points chosen for study included follicle growth and granulosa cell proliferation, based on the known proliferative effect of HH in cells of the Drosophila ovary [5–7] and in a variety of mammalian tissues and cell types [19]. In addition, effects of HH signaling on gene expression and steroid production by cultured granulosa cells were examined based on the reported effects of HH signaling to regulate differentiation in a variety of tissues [20–22].

MATERIALS AND METHODS

Materials

Random hexamer and deoxynucleotide triphosphates were obtained from Amersham (Piscataway, NJ), avian myeloblastosis virus reverse transcriptase (AMV-RT), and the Access RT-PCR kit from Promega (Madison, WI), and Taq polymerase from Fisher Scientific (Fair Lawn, NJ). Real-time RT-PCR reagents were obtained from Applied Biosystems (Foster City, CA). Primary antibodies for HH pathway proteins were purchased from Santa Cruz Biotechnology (Santa Cruz, CA): rabbit anti-human SHH, sc-9024, a pan HH antibody that binds all three HH ligands; goat anti-mouse PTCH1, sc-6047; and rabbit anti-human SMO, sc-13943. Biotinylated secondary antibodies and HRP-conjugated secondary antibodies were purchased from Jackson ImmunoResearch (West Grove, PA). Horseradish peroxidase (HRP)-conjugated streptavidin, Alexa488-conjugated goat anti-mouse IgG, propidium iodide (PI), and biotinylated-tyramide amplification kits (TSA Kit #21) were from Molecular Probes (Eugene, OR), and Nova Red substrate was from Vector Laboratories (Burlingame, CA). Mouse anti-human Ki67 was from Novocastra Laboratories (New Castle upon Tyne, UK) and mouse anti-bromodeoxyuridine (BrdU) (clone 3D4) was from BD PharMingen (San Diego, CA). Recombinant mouse SHH was purchased from R&D Systems (Minneapolis, MN), and cyclopamine and tomatidine from BioMol (Plymouth Meeting, PA). Culture media and media supplements were obtained from Life Technologies, Inc. (Grand Island, NY). Testosterone and BrdU were obtained from Sigma-Aldrich (St. Louis, MO). Ovine FSH (NIDDK-oFSH-20) was provided by Dr. A. F. Parlow, National Hormone & Peptide Program, Harbor-UCLA Medical Center (Torrance, CA). Ultra Low Attachment 96-well culture plates were obtained from Corning Costar (Corning, NY), Titertek 8-well Chamber Slides were from Nalge Nunc International (Rochester, NY), and Falcon 96-well and 6-well culture dishes were from Becton-Dickinson (Franklin Lakes, NJ).

Animals

Procedures were approved by the Cornell University Institutional Animal Care and Use Committee and are in accordance with the Guide for Care and Use of Laboratory Animals (National Academy of Science). CD-1 mice were purchased from Charles River Laboratories (Boston, MA) and housed in a colony with a 14L:10D cycle. Ovaries, testes, and duodena were collected from adult mice at least 42 days old, and ovaries were collected from immature mice on the day of birth or at 4, 7, 12, or 25 days of age. Uteri were obtained from pregnant mice 4 days after observation of a vaginal plug. Female mice that had been mated with vasectomized males were euthanized on Day 4 of pseudopregnancy, and corpora lutea were isolated by dissection with fine forceps. Ovarian surface epithelial cells were removed from the ovary by enzyme digestion prior to dissection of corpora lutea. Granulosa cells were obtained from 23-day-old mice 48 h after s.c. injection of 5 IU eCG. Large follicles were punctured with a 26-gauge needle and granulosa cells expressed into Dulbeccos Modified Eagle Medium, Hams F-12 Nutrient Mixture (DMEM-F12) and collected by centrifugation. Cells were resuspended in DMEM-F12, and any pieces of tissue, which might contain theca or interstitial cells, were removed by unit gravity sedimentation. Suspended granulosa cells were centrifuged and resuspended appropriately for the intended use. Cultured granulosa cells were uniform in appearance, indicating essentially pure preparations. Tissues and cells were either snap frozen in liquid nitrogen and stored at –80°C until preparation of RNA or processed for immunohistochemistry or culture as described.

RT-PCR

Nonquantitative RT-PCR assays were used to determine whether ovarian tissue expresses components of the HH pathway. Total RNA was isolated from ovaries, granulosa cells, and corpora lutea pooled from different mice [23]; three separate preparations were produced for each tissue. Single RNA preparations from Day 4 gestation uterus, duodenum, and adult testes were used as controls. RNA (7 µg) was reverse transcribed using AMV-RT and random hexamer primer. One preparation of each tissue was reverse transcribed at the same time, and this was repeated for each of the three tissue preparations. Aliquots of the cDNA (derived from 0.5 µg RNA) were used for multiple PCR reactions to amplify each of the HH pathway genes listed in Table 1 and peptidylprolyl isomerase A (Ppia, also known as cyclophilin), such that all PCR reactions except for Dhh (see below) were performed from a single RT reaction for each sample. Amplification consisted of a preincubation at 95°C for 5 min before adding Taq polymerase and then 40–45 cycles at 95°C for 30 sec, 52°–58°C for 30 sec (Table 1), and 72°C for 30 sec. Because genes encoding HH ligands share regions of similar nucleotide sequence, primers were designed to unique regions of each gene. All primer pairs span the position of one to three introns. Thus, the relatively short products expected from amplification of cDNA (210 to 524 bp; Table 1) could be distinguished from potential products amplified from contaminating genomic DNA which would be 2335 to 16 493 bp larger. For each set of primers, reactions using granulosa cell RNA as template were run with and without AMV-RT to confirm that the products obtained were not derived from laboratory contamination. Dhh was assayed using the Access RT-PCR kit with 0.5 µg RNA according to the manufacturer's recommendations. Representative RT-PCR products for each gene, generated using RNA from whole ovaries as template, were sequenced at the Cornell University DNA Sequencing Facility. RT-PCR products for Shh, generated using RNA from granulosa cells, corpora lutea, and whole ovary as templates, and RT-PCR products for Dhh, generated using RNA from granulosa cells, corpora lutea, uteri, and testes were also sequenced. Sequences obtained were compared to the refereed sequences from the National Library of Medicine.


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  		TABLE 1. Primers and annealing temperatures used in RT-PCR of hh pathway genes.

Quantitative real-time PCR assays were used to measure changes in expression of Gli1 in response to treatments and to measure mRNA levels of the three HH ligands in ovaries at different stages of development. RNA was prepared as previously described. Reverse transcription was performed using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). PCR was performed on cDNA from 0.05 µg RNA using the following commercially available mouse-specific Taqman Gene Expression Assays (Applied Biosystems): Gli1, Mm9949645_m1; Ihh, Mm00439613_m1; Dhh, Mm0043282C_g1; Shh, Mm00436527_m1. All primer pairs span the position of introns, preventing detection of genomic DNA, and utilize sequence-specific fluorescently labeled probes for detection of the PCR product. PCR was performed using an Applied Biosystems ABI 7000 Sequence Detection System. Standards were prepared from pooled RNA of Day 25 and adult ovaries. All samples for each gene were analyzed in a single assay. Results were standardized against concentrations of 18S in the same reverse transcription reactions as determined by PCR (Kit # 4319413E).

Immunohistochemistry

Tissues were fixed in 2% paraformaldehyde for 4 h at 4°C and embedded in paraffin. Next, 4-µm sections were deparaffinized in xylene, rehydrated, rinsed in 0.03% Triton X-100 in PBS followed by PBS. Antigen retrieval was performed by placing the slides in boiling 10 mM citrate buffer for 10 min. Endogenous peroxidase activity was quenched in 2% hydrogen peroxide in PBS. Nonspecific binding was blocked at room temperature for 30 min in PBS-1% BSA (PTCH1 and SMO) or 1% normal goat serum in PBS (pan HH). Sections were incubated with primary antibody (1 µg/ml) in PBS-1% BSA for 1 h at 37°C. For pan HH and PTCH1, slides were washed with PBS and incubated with 0.05 µg/ml biotinylated secondary antibodies (goat anti-rabbit IgG and donkey anti-goat IgG, respectively) in PBS-1% BSA for 1 h at 37°C followed by incubation in 0.25 µg/ml HRP-conjugated streptavidin for 20 min at room temperature. Slides were exposed to Nova Red solution for 5 min and counter-stained with Gills hematoxylin. Staining for SMO was intensified by tyramide amplification according to the manufacturer's instructions. Binding was detected with HRP-conjugated streptavidin, Nova Red, and counterstaining as previously described.

Staining for pan HH, PTCH1, and SMO were performed on ovaries from three immature mice (25 days old) and three adult mice (at least 42 days old). Representative sections from each ovary were scored for the intensity of staining in granulosa and theca cell layers in follicles at various stages of development from type 3 to type 8 (classified as described [24]). Briefly, type 2 are resting primordial follicles, type 3 are primary follicles with a single layer of granulosa cells, type 4 are secondary follicles with up to two layers of granulosa cells, type 5 have three or more layers of granulosa cells, type 6 follicles have begun to form an antral cavity, type 7 follicles have a fully formed antral cavity, and type 8 follicles are preovulatory. Relative intensities of staining were scored: –, no staining; +, moderate staining; or ++, strong staining. The morphology of type 2 (primordial follicles) was not adequate to allow staining in these follicles to be scored. Images were obtained using a Spot II Digital Camera (Diagnostic Instruments, Sterling Heights, MI). Images were obtained using identical light and exposure settings for each microscope lens. For each type of staining (pan HH, PTCH1, or SMO), identical brightness and contrast adjustments were applied to all images.

In order to confirm that the commercial antibodies used for the immunohistochemistry recognize the intended proteins, Western blots for pan HH, SMO, and PTCH1 were run. Tissue lysates were prepared in RIPA buffer (1.0% Nonidet P-40, 0.05% Na-deoxycholate, 0.1% SDS in PBS) containing freshly added proteinase inhibitors (100 µg/ml phenylmethylsulfonyl fluoride and 3 mg/ml aprotinin). Tissues were sonicated for 10 sec and lysates frozen until further analysis. Protein content of cell lysates was determined using the DC Protein Assay kit (Biorad, Hercules, CA). Protein lysates (200 µg/lane) were separated by 12% (pan HH and SMO) or 5% (PTCH1) SDS-polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride (PVDF) membranes. Membranes were blocked in tris-buffered saline (TBST; 20 mM Tris, pH 8.0, 150 mM NaCl, 0.05% Tween-20) containing 5% nonfat milk for 30 min at room temperature. Membranes were incubated at 4°C overnight in TBST-5% BSA containing 0.4 µg/ml of the appropriate antibody. Membranes were washed, incubated with 0.04 µg/ml HRP-conjugated secondary antibodies in TBST-5% nonfat milk for 30 min at room temperature, and washed. A chemiluminescent signal was generated using Western Blot Chemiluminescence Reagent (NEN, Boston, MA) and membranes were exposed to x-ray film (Kodak, Rochester, NY).

Granulosa Cell Culture, Proliferation Assays, and Gene Expression Analysis

Effects of increasing HH signaling were studied by treating granulosa cells with recombinant SHH, which binds both PTCH1 and PTCH2. Because SHH, IHH, and DHH appear to act similarly, recombinant SHH has been used in studies to activate the pathway in a variety of cells in vitro [25, 26] and in vivo [27]. Effects of decreasing HH signaling were studied by treating cells with cyclopamine, a plant steroidal alkaloid that inhibits the cellular response to HH signaling by antagonizing SMO [28–31]. Tomatadine, a compound that is structurally similar to cyclopamine but does not inhibit SMO, was used as a control. An initial dose-response experiment showed that SHH increased the percentage of granulosa cells that incorporated BrdU into DNA. For SHH at doses of 0, 0.1, 0.3, 1.0, and 3.0 µg/ml, the percentages of cells incorporating BrdU were 44.0%, 48.4%, 54.9%, 51.2%, and 52.6%, respectively. Based on this data, recommendations of the supplier, and previous studies using recombinant SHH [25, 26], a dose of 1 µg/ml was used in subsequent studies. In previous studies, doses of cyclopamine ranging from 1 to 10 µM [21, 26, 32] as well as higher doses (50 µM; [25]) were effective in inhibiting HH signaling. In the preliminary dose-response experiment, the percentage BrdU incorporation in granulosa cells treated with the tomatadine control at 1 µM (42.1%) and 2 µM (40.4%) were not substantially different from the media alone control (44.0%). The stimulatory effect of 1 µg/ml SHH on BrdU incorporation (51.2% in the presence of SHH vs. 44.0% in control) was blocked by 1 µM cyclopamine (41.3%) and 2 µM cyclopamine (39.7%). Based on this data, subsequent experiments were performed with 2 µM tomatadine and cyclopamine.

To study effects of altering HH signaling on proliferation, granulosa cells from eCG-primed mice were collected as described and cultured in chamber slides (2 x 105 cells/well) in DMEM-F12, supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, 0.25 µg/ml fungizone, 1 mM pyruvate, 2 mM glutamine, and containing 10% FBS (DMEM-F12–10% FBS). At 24 h, media was replaced with serum-free DMEM-F12 containing 100 ng/ml insulin, 5 µg/ml transferrin, 20 nM Na-selenite, and 0.1% BSA (DMEM-F12-ITS) and one of the following treatments: 1) no treatment, control; 2) 1 µg/ml SHH; 3) 2 µM cyclopamine; 4) 1 µg/ml SHH and 2 µM cyclopamine; and 5) 2 µM tomatidine. Following 2 days of treatment, cells were fixed in acetone and stained for Ki67 using mouse anti-human Ki67 antibody and PI as previously described [33]. Proliferation was further examined by adding 10 µM BrdU to some cultures during the last 24 h of culture, fixing cells, and staining with an anti-BrdU antibody and PI as previously described [33]. For each treatment in each experiment, images were obtained of four randomly chosen fields using a Spot II Digital Camera, and Ki67-positive and total cells (PI-stained nuclei), or BrdU-positive and total cells, were counted by two independent observers. Experiments were repeated three times each for Ki67 and BrdU, and each experiment included all treatments.

To study effects of HH signaling on levels of mRNA for the transcriptional target gene, Gli1, granulosa cells from eCG-primed mice were cultured in 6-well plates (1 x 106 cells/well) in DMEM-F12–10% FBS. At 24 h, media was replaced with DMEM-F12-ITS containing one of the following treatments: 1) no treatment, control; 2) 1 µg/ml SHH; 3) 2 µM cyclopamine. At 48 h, cells were removed from the wells with trypsin, rinsed, pelleted, and the pellets frozen for subsequent preparation of RNA and analysis by quantitative real-time RT-PCR. The experiment was repeated three times with separate granulosa cell preparations.

Measurement of Progesterone and E2 Secretion by Granulosa Cells

Granulosa cells were obtained from eCG-primed mice as described and cultured in 96-well plates (5 x 104 cells/well) in DMEM-F12–10% FBS containing 144 ng/ml testosterone (as substrate for aromatization to E2) and the following treatments: 1) no treatment, control; 2) 1 µg/ml SHH; 3) 2 µM cyclopamine; 4) 1 µg/ml SHH and 2 µM cyclopamine; and 5) 2 µM tomatidine. After 1, 2, and 3 days of culture, media were collected from the wells, centrifuged at 400 x g for 5 min to remove any cells, and the supernatant frozen at –20°C for later analyses. After the first day of culture, media was replaced daily with serum-free DMEM-F12-ITS containing testosterone and the same treatments as previously outlined. Media samples were assayed for progesterone and E2 using commercial RIA kits (Diagnostic Products, Los Angeles, CA) according to the manufacturer's instructions. Experiments were repeated four times, and each experiment included all treatments.

Whole Follicle Culture and Growth Assays

Secondary follicles ranging in diameter from 115 to 155 µm were dissected from 17- to 18-day-old mice and cultured according to the method of Fehrenbach et al. [34] with minor variations. Follicles of this size contain two layers of granulosa cells. Follicles were cultured in Corning-Costar Ultra Low Attachment 96-well plates (1 follicle/well) in DMEM-F12 containing 50 ng/ml ovine FSH and 50 µg/ml ascorbic acid. The following treatments were added to media at the beginning of culture: 1) no treatment, control; 2) 1 µg/ml recombinant SHH; 3) 2 µM cyclopamine; 4) 1 µg/ml SHH and 2 µM cyclopamine; and 5) 2 µM tomatidine. Media were replaced with fresh media containing the same treatments on Day 2 of culture. The diameter of each follicle (mean of horizontal and vertical diameters) was measured in digital images taken of each follicle on Days 0, 2, and 4 of culture. Image pixel length was calibrated to a hemacytometer grid. A total of 126 follicles were successfully cultured in five separate experiments (2 mice/experiment); 42 follicles were lost due to handling and to degradation leading to loss of the oocyte. Each experiment included all treatments.

On Day 4, follicles were fixed in 2% paraformaldehyde for 30 min at 4°C and stored in PBS-0.2% sodium azide at 4°C until staining for Ki67. Follicles were permeablized by incubation in PBS-2% BSA-0.1% Triton X-100 for 1 h at room temperature. This same buffer was used for antibody dilutions and washes. Follicles were incubated in 2 µg/ml mouse anti-Ki67 antibody overnight at room temperature, rinsed, and incubated in 0.25 µg/ml Alexa488-conjugated goat antimouse IgG overnight at room temperature. Follicles were rinsed and incubated in 0.25 µg/ml PI, a nuclear stain, and 30 µg/ml RNaseA at 4°C overnight. Follicles were rinsed in PBS, mounted on slides, and digital images taken with a Leica SP2 confocal microscope. Ki67-positive granulosa cells and total granulosa cells (PI-stained nuclei) were counted in three planes by two independent observers.

Statistical Analysis

Follicular growth data and E2 and progesterone production data were analyzed by a two-way randomized complete block ANOVA. Granulosa cell proliferation data and granulosa cell Gli1 mRNA data were analyzed by a one-way randomized complete block ANOVA. Quantitative RT-PCR analysis of Ihh, Dhh, and Shh were analyzed by a completely randomized (one-way) ANOVA. The Student-Newman-Keuls method was used for comparison of means when overall significance was observed.

RESULTS

Expression of Components of the HH Pathway in the Ovary

RT-PCR was used to determine whether components of the HH signaling pathway are expressed in the mouse ovary. The three HH ligands, Ihh, Shh and Dhh, are expressed in immature and adult ovaries, in granulosa cells isolated from 23-day-old eCG-primed mice, and in corpora lutea from pseudopregnant mice (Fig. 1). The two receptors, Ptch1 and Ptch2, as well as the mediator of HH signaling, Smo, are also expressed in all ovarian tissues examined. In addition, two targets and mediators of HH signaling, the transcription factor, Gli1, and Hhip are expressed in all ovarian tissues. Bands of the correct size were observed in all reactions performed in the presence of RT but were lacking in reactions performed without RT, indicating that the bands observed were generated from mRNA. Additional faint bands of larger transcripts were detectable in reactions for Ihh and Gli1 in the absence of RT and likely represent nonspecific products generated by contaminating genomic DNA. RT-PCR products were verified by cloning and sequencing products for each gene shown in Figure 1, generated using total ovarian RNA as template. In addition, the following RT-PCR products were confirmed by direct sequencing of the PCR reactions: Shh, generated using RNA from granulosa cells and corpora lutea; and Dhh, generated using RNA from granulosa cells, corpora lutea, uteri, and testes. All of the sequences obtained were ≥99% identical to refereed sequences from the National Library of Medicine. As expected based on previous studies, components of the HH pathway are also expressed in uterus, testis, and duodenum.


Figure 01
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  		FIG. 1. Expression of genes within the HH signaling pathway in ovarian and control tissues. Left panel: An ethidium bromide-stained gel of a single RT-PCR assay is shown for each gene, and is representative of results obtained in three separate assays. In each assay, a single RT reaction was prepared for each tissue and was used for multiple individual PCR reactions with primers specific for genes within the HH pathway. Assays for Dhh were performed separately (described in Materials and Methods). Right panel: Ethidium bromide-stained gels of RT-PCR reactions for each gene, in which AMV-RT was either included (+RT) or not included (–RT) in the reverse transcriptase reaction. GC, granulosa cells; CL, corpora lutea; Duod, duodenum; H20, water blanks in which template was omitted from PCR reactions; Ppia, run as a control for amount and quality of RNA used in the reactions.

Expression of Ihh, Dhh, and Shh mRNA during Ovarian Development

Because of the importance of the HH pathway in developmental processes, quantitative expression of the three HH ligands was examined in ovaries at different times during ovarian development (Fig. 2). Ihh and Dhh expression was low on the day of birth (Day 0), higher on Day 4 (P < 0.05), and remained elevated on all subsequent days (P < 0.05 vs. Day 0). On Day 25, when expression was highest, Ihh mRNA was 21-fold higher than on Day 0, and Dhh mRNA was 8-fold higher than on Day 0. In contrast, Shh mRNA expression was elevated on Day 0, and also on Days 7, 12, and 25. Shh expression was significantly lower on Day 4 and in adult ovaries.


Figure 02
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  		FIG. 2. Expression of HH ligands, Ihh, Dhh, and Shh, in ovarian RNA obtained at different stages of development, as determined by quantitative real-time RT-PCR. Day 0 = day of birth. Three samples were analyzed at each time. Bars represent the mean ± SEM of results obtained using three separate RNA preparations at each time point. Bars with different superscripts are significantly different (P < 0.05).

Localization of HH Family Proteins by Immunohistochemistry

Staining for HH ligands using a pan HH antibody is most pronounced in the granulosa cells of antral follicles (type 6, 7, and 8; Table 2 and Fig. 3, a and e). In preantral follicles (type 5; Fig. 3f) and small follicles (types 3 and 4; Fig. 3g), the staining for pan HH is more variable. In all follicles, staining for pan HH is generally less intense in theca cells than in granulosa cells (Table 2). Staining for PTCH1 is strongest in granulosa cells and theca cells of antral follicles (Fig. 3, i and m), but is also moderate to strong in granulosa cells and theca of preantral and small follicles (Fig. 3, i, n, and o). In follicles of all types, staining for SMO is moderate to strong with little difference in intensity between granulosa cells and theca (Table 2; Fig. 3, q, u, v, and w). Corpora lutea stain prominently for pan HH, PTCH1, and SMO (Fig. 3, c, k, and s). Specificity of antibodies were verified using control tissues known to express HH pathway proteins including epididymis (pan HH, Fig. 3h) and uterus isolated on Day 4 of pregnancy (PTCH1, Fig. 3p and SMO, Fig. 3x).


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  		TABLE 2. Intensity of immunostaining for pan HH, PTCH1, and SMO in granulosa and theca cells of various classes of follicles.


Figure 03
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  		FIG. 3. Immunohistochemistry of HH pathway proteins in mouse ovaries. a–h) Representative staining for HH ligands using a pan HH antibody that detects SHH, IHH, and DHH. i–p) Staining for PTCH1. q–x) staining for SMO. In each set of panels, low power images of ovaries from 25-day-old mice and corpora lutea from adult mice are shown (a–d, i–l, and q–t), including sections incubated with IgG instead of antibodies (Ab–) to show nonspecific binding (b, d, j, l, r, and t). Each set of panels shows higher magnification images of type 7 antral follicles (e, m, u), type 5 preantral follicles (f, n, v), and type 3 primary follicles (g, o, w). Control tissues shown are epididymis stained for pan HH (h), and uterus from Day 4 of pregnancy stained for PTCH1 (p) and SMO (x).

Western immunoblotting was performed to assure the specificity of the antibodies used for immunohistochemistry (Fig. 4). Lysates prepared from whole ovaries (Day 25 and adult) and granulosa cells produced bands of the expected size: pan HH, ~21 kDa with lesser bands at ~25 kDa; SMO, 85 kDa; PTCH1, 165 kDa. The multiple bands in the pan HH Western blot are due to differences in n- and c-terminal lipid attachment during posttranslational modification [35].


Figure 04
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  		FIG. 4. Western blot analyses of pan HH, PTCH1, and SMO in ovarian and uterine protein lysates performed with the same antibodies used for immunohistochemistry. Blots incubated with antibody (left panels) or IgG (nonspecific binding, right panels) are shown. Molecular weights (in kDa) are indicated.

Effect of HH Signaling on Growth of Follicles In Vitro

The ability of HH signaling to alter follicle growth rate in vitro was tested using preantral follicles of 115 to 155 µm diameter that were isolated from ovaries of immature 17- to 18-day-old mice. Preantral follicles increased in diameter over 4 days of culture and the number of layers of granulosa cells increased in response to each treatment (Fig. 5, A and B). The percentages of follicles that grew greater than 25% between Day 0 and 4 of culture were 51.9% for controls, 90.3% for SHH, 45.8% for cyclopamine, 65.0% for SHH plus cyclopamine, and 62.5% for tomatadine. While the average diameters of follicles on Day 0 and Day 2 of culture did not differ among treatments, follicle diameter on Day 4 was increased by SHH (Fig. 5A; P < 0.05). Treatment with cyclopamine had no effect on follicle diameter but concomitant treatment with cyclopamine and SHH prevented the SHH-induced increase in diameter. Tomatidine, a compound that is structurally related to cyclopamine but does not inhibit SMO, had no effect. Representative follicles that either increased in diameter between Days 2 and 4 of culture or failed to grow, were analyzed for expression of the cell proliferation marker Ki67. There was a positive correlation between the increase in follicle diameter and the percentage of Ki67-positive cells (Fig. 5, C and D).


Figure 05
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  		FIG. 5. Effect of HH signaling on growth of preantral follicles in vitro. A) Preantral follicles were cultured for four days in control media (n = 27) or media containing 1 µg/ml SHH (n = 31), 2 µM cyclopamine, a pharmacological inhibitor of the HH pathway (n = 24), 2 µM tomatidine (n = 24), a compound structurally related to cyclopamine but unable to inhibit the pathway, or 2 µM cyclopamine and 1 µg/ml SHH (n = 20). Follicle diameters were measured on Days 0, 2, and 4 of treatment using digital images of individual follicles. *P < 0.05 vs. other treatments on the same day of culture. B) Phase-contrast images of a representative follicle over four days of culture showing an increase in diameter and an increase in the number of the layers of granulosa cells. C) Correlation between the percent growth of preantral follicles from Days 2 to 4 of culture and the percentage of granulosa cells expressing Ki67. Representative follicles that grew or did not grow during culture were costained for Ki67 and PI, and the percent Ki67-positive cells determined. D) Confocal images of Ki67 staining of representative follicles that grew (left panel) or did not grow (right panel) from Days 2 to 4 of culture. Nuclei positive for Ki67 and PI appear bright white, while nuclei positive for PI alone appear grey.

Effect of HH Signaling on Granulosa Cell Proliferation and Gene Expression In Vitro

Treatment of granulosa cells isolated from eCG-primed mice with recombinant SHH induced an 89.9% increase in expression of Ki67 relative to controls (P < 0.05, Fig. 6A). SHH also induced a 30.0% increase in incorporation of BrdU into DNA relative to controls (P < 0.05, Fig. 6B). Treatment with cyclopamine alone had no effect on expression of Ki67 or incorporation of BrdU, but concomitant treatment with cyclopamine and SHH prevented the effects of SHH. Treatment with the control compound, tomatidine, had no effect on expression of Ki67 or incorporation of BrdU (Fig. 6, A and B).


Figure 06
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  		FIG. 6. Effect of HH signaling on proliferation of granulosa cells in vitro. Granulosa cells from 23-day-old eCG-primed mice were cultured in control media or media containing 1 µg/ml SHH, 2 µM cyclopamine, a pharmacological inhibitor of the HH pathway, 2 µM tomatidine, a compound structurally related to cyclopamine but unable to inhibit the pathway, or 2 µM cyclopamine and 1 µg/ml SHH. A) Percentage of granulosa cells expressing Ki67. B) Percentage of granulosa cells incorporating BrdU. Bars represent the mean ± SEM of results obtained in three experiments using separate granulosa cell preparations. *P < 0.05 vs. other treatments within each panel.

Effects of HH signaling on the downstream transcriptional target gene, Gli1 were examined to determine if HH has a direct effect on granulosa cells. SHH increased expression of Gli1 mRNA 140% compared to control cultures (P < 0.05, Fig. 7), while treatment with cyclopamine reduced expression to 32% of control cultures (P < 0.05).


Figure 07
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  		FIG. 7. Effect of HH signaling on expression of Gli1 mRNA by granulosa cells in vitro. Granulosa cells from 23-day-old eCG-primed mice were cultured in control media, media containing 1 µg/ml SHH, or media containing 2 µM cyclopamine. After 24 h, cells were collected and RNA prepared. Gli1 mRNA concentrations were determined by quantitative real-time RT-PCR. Bars represent the mean ± SEM of results obtained in three experiments using separate granulosa cell preparations. Bars with different superscripts are significantly different (P < 0.05).

Effect of HH Signaling on Granulosa Cell Steroidogenesis

Production of progesterone and E2 by cultured granulosa cells from eCG-primed mice were determined in response to treatments to stimulate or inhibit the HH pathway. Treatment with SHH did not alter production of progesterone compared to that in control cultures or in cultures treated with tomatidine (Fig. 8). In contrast, cyclopamine increased progesterone production on Day 3 of culture 50% relative to controls. Progesterone production was similarly elevated in response to treatment with cyclopamine in the presence of SHH on Day 3 of culture (Fig. 8). None of the treatments altered E2 production (Fig. 8). In all cultures, production of E2 was highest on the first day of culture and decreased on Days 2 and 3.


Figure 08
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  		FIG. 8. Effect of HH signaling on granulosa cell steroidogenesis. Granulosa cells from 23-day-old eCG-primed mice were cultured in control media or media containing 1 µg/ml SHH, 2 µM cyclopamine, a pharmacological inhibitor of the HH pathway, 2 µM tomatidine, a compound structurally related to cyclopamine but unable to inhibit the pathway, or 2 µM cyclopamine and 1 µg/ml SHH. The concentrations of progesterone and estradiol in the culture media were determined. Bars represent the mean ± SEM of results obtained in four experiments using separate granulosa cell preparations. *P < 0.05 vs. other treatments on the same day of culture.

DISCUSSION

Our findings indicate that major components of the HH pathway are expressed in the mouse ovary and that granulosa cells are one of the targets of HH signaling in the follicle. RT-PCR analyses followed by DNA sequencing demonstrated the presence of mRNA for the ligands (Ihh, Shh and Dhh), the receptors (Ptch1 and Ptch2), the signal transducer (Smo), downstream transcription factor (Gli1), and pathway regulator (Hhip). Transcripts for these genes were detected in each of the ovarian tissues studied; immature and adult ovaries, granulosa cells, and corpora lutea. As expected based on previous studies, transcripts were also detected in uterus, testis, and duodenum, tissues analyzed as controls [27, 36–38]. Immunohistochemistry of HH ligands (using a pan HH antibody), and PTCH1 and SMO proteins are consistent with the mRNA expression data. Specific staining was detectable in granulosa and theca cells of a wide range of follicles and was most intense in antral follicles. HH, PTCH1, and SMO proteins were all detectable in corpora lutea.

Analysis of ovarian expression of the HH ligands during development showed low levels of Ihh and Dhh expression on the day of birth, increases in both on Day 4, and maintenance of elevated levels in prepubertal and adult mice. The data are consistent with detection of HH protein in growing follicles by immunohistochemistry and with a previous report which showed that Ihh and Dhh mRNA are expressed in the granulosa cells of primary through antral follicles [4]. In the mouse, primordial follicle formation has not yet occurred at the time of birth but occurs within the first several days of life. By Day 4, some primordial follicles have begun to grow and develop to the primary stage and these follicles may contribute to the first increases in ovarian Ihh and Dhh expression observed. The pattern of Shh expression differed from that of Ihh and Dhh in that it was highest in ovaries on the day of birth, decreased transiently on Day 4, returned to the higher level of expression in prepubertal mice and declined in adults. The biological significance of fluctuations in Shh expression remains to be determined but is of interest because important processes such as the formation of primordial follicles and the first wave of follicle growth occur during this developmental time frame. A previous study reported that expression of Shh was not detectable in the mouse ovary by in situ hybridization or RT-PCR [4]. This might be explained by lower relative levels of expression of Shh compared to Ihh and Dhh and the developmental time points chosen for study.

A goal of the current study was to determine whether the HH pathway in the ovary is functional. We first investigated a potential role in regulating the proliferation of granulosa cells because in the Drosophila ovary HH signaling increases proliferation of somatic stem cells, which generate the layer of epithelial cells that encapsulate germ cell cysts [7]. In mammals, HH signaling regulates cell proliferation in numerous tissues (reviewed in [1]) by promoting transcription of cyclin D and cyclin E [39] and blocking cell cycle arrest by the p21 cip1/waf1 cell cycle inhibitor [19, 40]. In the current study, treatment of preantral follicles with SHH in vitro increased growth relative to controls and growth was associated with proliferation of granulosa cells. Treatment with cyclopamine, a plant alkaloid that has been shown to inhibit SMO, did not alter follicle growth, suggesting that the basal rate of follicle growth under the culture conditions used is not dependent on HH signaling. The fact that treatment with cyclopamine in the presence of SHH prevented the SHH-induced increase in growth indicates that the concentration of cyclopamine used in these experiments was adequate to prevent HH signaling. Treatment of granulosa cells isolated from preovulatory follicles of eCG-primed mice with SHH in vitro increased two measures of cell proliferation: expression of Ki67 and incorporation of BrdU, while treatment with cyclopamine had no effect. These results suggest that increased signaling through HH may induce a small increase in granulosa cell proliferation, but HH signaling is not required to maintain basal proliferation under the culture conditions used. Cellular responses to HH signaling are tightly regulated. For example, HH increases transcription of Ptch1 mRNA while increased PTCH1 protein subsequently attenuates signaling through SMO [11, 13–15]. HH also increases transcription of Hhip and HHIP protein binds to HH ligands and prevents interaction with PTCH [11, 17, 18]. Cancers in a wide array of tissues result when HH signaling is abnormally activated by mutations in PTCH or SMO or overactivation of HH ligands [3, 11, 12]. Thus, a limited proliferative response to HH signaling might be expected in nontransformed cells.

Effects of SHH on steroid production in vitro was examined as an additional measure of whether granulosa cells may be responsive to HH signaling. In addition to regulating cell proliferation, HH directs cell fate in the Drosophila ovary [5, 6] and the differentiation of cells in a number of mammalian tissues. For example, in cartilage [20], prostatic epithelium [21], and dental epithelium [22], HH signaling stimulates proliferation and also affects differentiation. Production of E2 and progesterone by granulosa cells change as they differentiate during follicle development (reviewed in [41]); granulosa cells acquire LH receptors and produce increased levels of E2 in response to FSH and LH. After exposure to the preovulatory LH surge, steroid production shifts; production of E2 declines while production of progesterone increases as granulosa cells differentiate into luteal cells. When granulosa cells are cultured, they spontaneously acquire some of the characteristics of luteal cells, producing progressively lower amounts of E2 and increased levels of progesterone over time. In the current study, production of E2 by granulosa cells from eCG-treated mice declined during culture, while production of progesterone increased, consistent with cell culture-induced luteinization. While treatment with SHH had no effect, blocking HH signaling with cyclopamine increased progesterone production. Production of E2 was not affected by treatments. These results are consistent with a potential role of HH signaling to alter functions of granulosa cells associated with differentiation, such as steroid production. Data from Wijgerde et al. [4] suggest that signaling through the HH pathway may decrease in preovulatory follicles after exposure to the LH surge, based on dramatically decreased expression of Ihh in granulosa cells and Ptch1 in theca cells. In addition, microarray analysis of cumulus oocyte complexes indicates that Ihh and Dhh expression decrease following the LH surge [42]. An interesting possibility, consistent with the observed stimulatory effect of cyclopamine on progesterone production in vitro, is that a decline in HH signaling after the LH surge may promote the differentiation of granulosa cells into luteal cells.

The effects of SHH on granulosa cell proliferation and progesterone production in vitro identify granulosa cells as a potential target for reception of HH signaling in the follicle. This interpretation is consistent with our findings that granulosa cells from eCG-treated mice express Ptch1, Ptch2, Smo, Gli1, and Hhip mRNA, transcripts expected in cells with a functional HH pathway. Granulosa cells from preantral and antral follicles also express PTCH1 and SMO proteins as well as HH ligand. The fact that treatment of granulosa cells with SHH increased levels of mRNA for Gli1, a transcriptional target of HH signaling, while treatment with cyclopamine decreased Gli1 mRNA also indicates that granulosa cells respond to HH signaling. Other examples of cell types that both synthesize a HH ligand and respond to HH signaling have been identified. For example, in the developing tooth, SHH that is synthesized in the dental epithelium acts as an autocrine mitogen in the epithelium and a paracrine mitogen for underlying mesenchymal cells. Autocrine HH signaling has also been reported in the embryonic skin [43] and developing lung [44]. Thus, substantial evidence exists for autocrine signaling through the HH pathway. Paracrine signaling through the HH pathway also occurs in many tissues [1]. Wijgerde et al. reported that levels of Ptch1 and Gli1 mRNA, detected by in situ hybridization of mouse ovaries, were highest in theca while levels of Ptch1 and Gli1 in the granulosa layer were not prominent, but appeared to be above background levels [4]. An interpretation of the data offered by the authors is that HH ligand produced in granulosa cells may signal to the adjacent theca, leading to increased expression of HH target genes. This possibility is supported by studies in a number of cell types in which HH signaling has been shown to increase levels of mRNA for target genes including Ptch1, Gli1, and Hhip [13–17]. In addition, it is consistent with the many examples of paracrine signaling between epithelial cells that secrete HH ligand and adjacent mesenchymal cells that respond to the signal [1]. Recent studies, however, suggest that another potential role for increased expression of Ptch1 in HH-target cells is to confine the field of responding cells to within a limited distance from the source of HH secretion [14, 15]. According to this model, once PTCH1 protein is expressed, it sequesters HH, preventing its action beyond the target field. Perhaps one consequence of increased Ptch1 mRNA expression in the theca is to prevent diffusion of HH ligand to neighboring follicles or interstitial cells. Elevated levels of Ptch1 mRNA in theca relative to granulosa cells may not necessarily indicate that HH signaling occurs in theca cells rather than in granulosa cells. Instead, it may indicate that theca cells define the limit of HH signaling in the follicle.

Because deletion of genes within the HH pathway in mice has generally resulted in early embryonic mortality [16, 20, 45–48], these experiments have not provided information about the role of HH signaling in ovarian function. An exception is the Dhh null mouse, in which males are infertile due to defects in the development of seminiferous tubules, while females are fertile [37]. Early studies failed to detect Dhh expression in the ovary and this was suggested as an explanation for the lack of a detectable defect in female Dhh null mice [37]. Taken together with our results and recent results of others, which provide evidence for expression of Dhh in the ovary [4, 42], DHH is apparently not essential for ovarian function. It is possible that IHH or SHH, which are also expressed in the ovary, may compensate for lack of DHH in Dhh null mice. Future studies using transgenic mice, in which genes that function within the HH signaling pathway are conditionally deleted in the ovary, will be important to determine the physiological role of the HH pathway.

The data presented here demonstrate that components of the HH pathway are expressed in the mouse ovary and show for the first time that HH signaling is functional in granulosa cells. These findings provide a basis for future investigations to define cell-specific responses to HH signaling in the follicle and to determine how the pathway modulates follicle development.

FOOTNOTES

1Supported by NIH grant HD 32535. Back

Correspondence: 2Susan M. Quirk, Department of Animal Science, Morrison Hall, Cornell University, Ithaca, NY 14853. FAX: 607 255 9829; e-mail: smq1{at}cornell.edu

Received: 8 May 2006.

First decision: 31 May 2006.

Accepted: 22 March 2007.

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Hedgehog signalling in endocrine development and disease
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