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BOR - Papers in Press, published online ahead of print August 16, 2006.
Biol Reprod 2006, 10.1095/biolreprod.105.050344
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BIOLOGY OF REPRODUCTION 75, 734–740 (2006)
DOI: 10.1095/biolreprod.105.050344
© 2006 by the Society for the Study of Reproduction, Inc.

Research Article

Granulosa Cells Promote Differentiation of Cortical Stromal Cells into Theca Cells in the Bovine Ovary1

Makoto Orisaka 3, Kimihisa Tajima 3, Tetsuya Mizutani 45 , Kaoru Miyamoto 45 , Benjamin K. Tsang 67, Shin Fukuda 3, Yoshio Yoshida 3, and Fumikazu Kotsuji 2 3

Departments of Obstetrics and Gynecology3 and Biochemistry,4 University of Fukui, Fukui 910-1193, Japan CREST,5 JST (Japan Science and Technology Agency), Saitama 322-0012, Japan Departments of Obstetrics & Gynecology6 and Cellular & Molecular Medicine,7 University of Ottawa, Ottawa Health Research Institute, Ottawa, Ontario, Canada K1Y 4E9

ABSTRACT

Formation of a theca cell (TC) layer is an important physiologic event that occurs during early follicular development. Nevertheless, little is known concerning the nature and regulation of the formation of the TC layer during follicular growth. Using an established coculture system in this study, we examined the hypothesis that stromal cells differentiate into TCs during early follicular development and that this process involves interaction with granulosa cells (GCs). Ovarian stromal cells from the bovine ovarian cortex (SC) and medulla (SM) were cultured with or without GCs from small antral follicles. The presence of GCs increased the number of lipid droplets and mitochondria, and it stimulated androstenedione production in SC and SM. However, luteinizing hormone/choriogonadotropin receptor (LHCGR) mRNA abundance and hCG-induced cAMP and androstenedione production were increased in SC but not in SM by the presence of GCs. The present results indicate that GCs are involved in the functional differentiation and the acquisition of LH responsiveness in stromal cells of the ovarian cortex. We suggest that GC-SC interaction is important in the formation of the TC layer during early follicular development, although the nature of this interaction remains to be determined.

androgen, follicle, granulosa cells, LH receptor, ovary, theca cells

INTRODUCTION

Although it is generally accepted that early follicular development is influenced by ovarian autocrine/paracrine regulators [1–7], our knowledge of the factors controlling follicle development from the primordial stage to the early antral stage is limited [2, 8]. The appearance of the theca cell (TC) layer is a key event for early follicular development, as evidenced by: 1) the coincidence between the organization of the TC layer and the responsiveness of bovine follicular growth and steroidogenesis to gonadotropins [9, 10]; 2) the provision by the TC layer of structural support and a blood supply containing ovarian regulators for the developing follicle [1, 11, 12]; and 3) production by TCs of aromatizable androgen for granulosa cell (GC) estrogen biosynthesis and stimulation of early follicular growth through androgenic products of TCs [13–18].

Using a GC-TC coculture system, we previously demonstrated the importance of TC-GC interaction in the control of follicular development [19–22]. Results of those studies showed that TCs maintained epitheliallike appearance and androgenic capacity when cocultured with GCs, but TCs became fibroblastic and produced less androgen when cultured alone [19], raising the possibility that GC-derived signals stimulate stromal cell differentiation into functional TCs. In the present study we assessed this hypothesis using the above-mentioned coculture system and demonstrated that GCs are involved in the functional differentiation and acquisition of LH responsiveness in stromal cells of the ovarian cortex. These observations support the notion that the ovarian stroma is the cellular origin of TCs and that GCs stimulate this differentiation process.

MATERIALS AND METHODS

Isolation of Ovarian Stromal Cells

Ovaries with extensive stromal tissue but without corpus luteum from adult cycling heifers were transported from a local abattoir in an ice-cold buffered salt solution. Ovarian stromal cells were isolated from the loose connective tissue of both the cortex and the medulla. Following removal of the ovarian surface epithelium using scissors, thin strips (approximately 2–3 mm thick) of stromal tissue containing either small numbers of preantral follicles (e.g., primordial, primary, and secondary follicles) from the ovarian cortex or small blood vessels from the ovarian medulla were dissected, respectively, for preparation of cortical stromal cells (SC) and medullary stromal cells (SM). Follicles and blood vessels then were removed with a scalpel under a dissecting microscope. The strips were cut into small fragments, and the cells were dissociated for 45 min at 37°C in Hanks-Hepes buffer containing collagenase (2150 U/ml, type 1; Sigma Chemical Co., St. Louis, MO), DNase (100 U/ml; Sigma Chemical Co.), bovine serum albumin (0.4% [v/v]; Sigma Chemical Co.), and glucose (0.2% [w/v]; pH 7.4) with continuous stirring at 800 rpm and then for 7 min in 0.25% (w/v) pancreatin (Sigma Chemical Co.) in a Hanks-Hepes buffer. Dispersed cells were filtered through Cell Strainer (80 µm; Becton Dickinson Labware, Franklin Lakes, NJ), washed three times in a culture medium consisting of Waymouth MB 752/1 medium (Invitrogen Corp., Carlsbad, CA), Hanks solution (Nissui Pharmaceutical Co. Ltd., Tokyo, Japan), and fetal calf serum (FCS; 6:3:1 [v/v/v]; Invitrogen Corp.) supplemented with streptomycin (100 µg/ml; Invitrogen Corp.) and penicillin (100 U/ml; Invitrogen Corp.). The cell viability, estimated using trypan blue exclusion, was 90%-95%.

Purity of Stromal Cells: Immunocytochemical Study

The purity of ovarian stromal cell preparations was evaluated immunocytochemically as described by Duleba et al. [23], with slight modifications. Briefly, 1000 stromal cells from the cortex and medulla of the ovaries (10 000 cells/mm3 in PBS) were placed onto poly-L-lysine-coated glass slides, fixed in 95% ethanol (pH 7.4; 18 h; room temperature), washed twice in PBS, treated with 1% H2O2 in methanol (10 min, to block endogenous peroxidase activity), and then washed in PBS. The slides were incubated (10 min) with Non-Specific Staining Blocking Reagent (DakoCytomation, Carpinteria, CA) to prevent nonspecific staining) and then with antivimentin (mouse; 1:20; BioGenex, San Ramon, CA), anticytokeratin AE1 + AE3 (mouse; 1:50; DakoCytomation), or anti-von Willebrand factor (mouse; 1:200; DakoCytomation) antibodies (3 h; room temperature in a humidified chamber), for the identification of mesenchymal, epithelial, and endothelial cells, respectively. Slides then were washed with PBS, incubated with biotinylated secondary antibody (10 min, 37°C), rinsed with PBS, and incubated with peroxidase-conjugated streptavidin (60 min, 37°C). Color reaction was initiated by incubating the sections for 10 min with diaminobenzidine-H2O2 (ENVISION/HRP [DAB]; DakoCytomation). The slides were counterstained with hematoxylin, dehydrated in gradient alcohol, and de-paraffined in xylene. The percentages of SC cells that were stained for vimentin, cytokeratin AE1 + AE3, and von Willebrand factor were 95.4% ± 3.4%, 0%, and 6.6% ± 1.8%, respectively, whereas those of SM cells were 92.3% ± 6.6%, 0%, and 10.8% ± 2.5%, respectively.

Preparations of Granulosa and Theca Cells

GCs and TCs were collected from small antral follicles (2–4 mm diameter), as described previously [19]. Only ovaries with a regressing corpus luteum were used [24]. Briefly, GCs were harvested from follicles using aseptic needle aspiration and were washed three times. Washed cells were filtered through Cell Strainer (80 µm) and resuspended in culture medium. Cell viability was estimated at 30%-42% using trypan blue exclusion testing. For TC preparation, follicles with clear surfaces were cut into halves, and the theca interna layer was removed with fine forceps. The GCs were removed by scraping with a scalpel under stereomicroscopy. The remaining TC layer was minced and dissociated into isolated cells by incubation with a Hanks-Hepes buffer containing collagenase, DNase, and bovine serum albumin, and subsequently with pancreatin in Hanks-Hepes buffer as described above for the preparation of stromal cells. Dispersed cells were filtered through Cell Strainer (80 µm) and washed three times. Cell viability, which was estimated by trypan blue exclusion, was 90%–95%.

Coculture on a Collagen Membrane

To examine the influence of GCs on ovarian stromal cell differentiation, a coculture of GCs and ovarian stromal cells was prepared on a collagen membrane as described previously for GC-TC coculture [19]. The double chambers were separated by a type 1 collagen membrane (thickness, 70 µm; area, 8 cm2; Koken Co. Ltd., Tokyo, Japan), which is permeable to molecules smaller than 12.5 kDa [19, 20]. The GCs (1 x 106 viable cells per dish) were cultured onto the membrane for 24 h in 1.5 ml culture medium consisting of Waymouth MB 752/1 medium, Hanks solution, and 10% FCS to allow for GC attachment (6:3:1 [v/v/v]), supplemented with streptomycin (100 µg/ml) and penicillin (100 U/ml). Cultures were carried out at 37°C under an atmosphere of 5% CO2 in humidified air. The membrane was then turned over and transferred into serum-free Ham F-12 medium (Nissui Pharmaceutical Co. Ltd.) supplemented with transferrin (10 µg/ml). Freshly prepared ovarian stromal cells (1 x 106 viable cells per dish) or TCs (1 x 106 viable cells per dish) were seeded on the opposite side and cultured. As controls, ovarian stromal cells or TCs were cultured alone on one side of a collagen membrane. For steroid assays and RT-PCR analyses, stromal cells or TCs were cultured in serum-free Ham F-12 medium for up to 48 h. The mean number of viable granulosa cells at the end of culture was 4.2 x 105, regardless of the type of cocultured cell. Cells for electron microscopic assessment were cultured in 10% FCS-containing medium because cell micro-organelles such as lipid droplets and secretory products were lost after 48 h culture under serum-free conditions.

Scanning and Transmission Electron Microscopy

After 48 h of culture under serum-supplemented conditions with or without GCs, stromal cells were fixed with 4% glutaraldehyde, postfixed in 1% osmium tetroxide, and then dehydrated in a series of ethanol baths. They were then coated with gold/palladium for scanning electron microscopic observation with an electron microscope (Hitachi S-450; Hitachi High-Technologies Corp., Tokyo, Japan). For transmission electron microscopy, the ovarian stromal cells were stained in 2% uranyl acetate solution before dehydration and were embedded in Araldite. Ultrathin sections were doubly stained in uranyl acetate in 90% methanol and lead citrate and were examined using an electron microscope (Hitachi H-7000; Hitachi High-Technologies Corp.). As indicators of steroidogenic potential, the numbers of lipid droplets and mitochondria in 90 SC cells and 60 SM cells were determined.

Steroid Assays

The spent media were collected from the apical and basal chambers and stored at –20°C pending steroid assays. Androstenedione levels in the culture media from the stromal side were measured using a double-antibody RIA (125I). All samples were analyzed in the same assays; the intraassay coefficient of variation was 6.0%.

Measurement of mRNA

Total cellular RNAs were extracted with TRIzol (Invitrogen Corp.) by the guanidium acid-isothiocyanate-phenol-chloroform method, in accordance with the manufacturer's protocol. Then, RT-PCR analyses for bovine 17alpha-hydroxylase/C17–20 lyase (CYP17A1), cholesterol side-chain cleavage cytochrome P450 (CYP11A1), 3beta-hydroxysteroid dehydrogenase (HSD3B1), luteinizing hormone/choriogonadotropin receptor (LHCGR), Steroidogenic acute regulatory protein (STAR), Steroidogenic Factor-1 (NR5A1), vascular endothelial growth factor (VEGF), and ARBP coding for acidic ribosomal phosphoprotein (as internal control) [25] were performed on total RNAs from freshly isolated cells (SC, SM, and TCs) or cultured cells following recovery by trypsin treatment, using the specific primers indicated in Table 1.


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  		TABLE 1. Sequences for target gene primers

The RNAs were reverse transcribed in 40 µl First-Strand Buffer (3 mM MgCl2, 75 mM KCl, 50 mM Tris-HCl, pH 8.3) containing 500 µM deoxynucleotide triphosphate (dNTP), 10 mM dithiothreitol, 200 units of SuperScript III RNase H free reverse transcriptase (Invitrogen Corp.), 200 ng random hexamers, and 2 µg total RNA. The target cDNAs of freshly prepared or cultured cells were amplified for 30 or 35 cycles (as detailed in the figure legends) at 94°C for 20 sec, 60°C for 30 sec, and 72°C for 60 sec, using dNTP (0.2 mM) and 1.5units of TaKaRa Ex Taq (TaKaRa Shuzo Co. Ltd., Kyoto, Japan) in a thermal cycler. Aliquots of PCR products were electrophoresed on 1.5% agarose gels and visualized with ethidium bromide staining. The relative integrated density of each band was scanned and digitized using FluorChem (Alpha Innotech Co., San Leandro, CA). The ratios of densitometric readings of the amplified target cDNA and ARBP were calculated.

For more quantitative assessment of LH receptor expression, real-time quantitative PCR analysis of LHCGR mRNA was performed on the cDNAs of stromal cells (SC or SM) and TCs, using a sequence detection instrument and software (ABI PRISM 7000; PE Applied Biosystems, Foster City, CA). The LHCGR primers used for amplification are indicated in Table 1. Amplification reaction was performed using Power SYBR Green PCR Master Mix kit (PE Applied Biosystems). The thermal conditions included an initial denaturation step at 95°C for 10 min and 35 cycles at 95°C for 15 sec, and 60°C for 1 min. The level of LHCGR mRNA was expressed as a ratio to the ARBP mRNA value.

Cyclic AMP Assays

A functional analysis of LH receptors in stromal cells was examined by measuring intracellular cAMP concentrations with a Cyclic AMP EIA Kit (Cayman Chemical, Ann Arbor, MI) after hCG challenge. Cultured stromal cells (SC or SM) were washed with warm medium and then preincubated in 1.5 ml serum-free medium in the presence of 1.0 M 3-isobutyl-1-methylxanthine (Sigma Chemical Co.) for 30 min at 37°C to prevent the breakdown of cAMP. After removal of the phosphodiesterase inhibitor, hCG (0.01–1 IU/ml; National Hormone & Peptide Program, Harbor-UCLA Medical Center; Torrance, CA) was added to the cells. Incubation then was continued for 30 min at 37°C. After incubation the medium was removed, and the cells were extracted with 1.0 ml of 0.1 M HCl. Intracellular cAMP levels were determined after acetylation procedure in accordance with the manufacturer's instructions.

Statistical Analysis

Each study was repeated three or four times. Data were assessed using one-way or two-way ANOVA, except for use of the unpaired t-test for electron microscopic data and real-time PCR analysis with a statistical analysis program (Prism 4.03 statistical software; GraphPad Software Inc., San Diego, CA). Differences between treatment groups were analyzed using Tukey or Bonferroni post-test; significance was inferred for P < 0.05.

RESULTS

Electron Microscopy of Cultured Ovarian Stromal Cells

To study the effect of GCs on ovarian stromal cells, isolated SC and SM were cultured with or without GCs in serum-supplemented medium for 48 h. The morphology of the cultured stromal cells was examined using an electron microscope. Figure 1 shows that SC cultured alone were thin, flat, and spindle shaped under scanning electron microscopy (Fig. 1A). In contrast, the apical surface of SC cocultured with GCs appeared to be more convex but also had a spindle shape (Fig. 1B). Scattered secretory products were observed on the cellular surface. Numerous filopodia were spread over the cell surfaces and connected to adjacent cells (Fig. 1B). In addition, transmission electron microscopy revealed that coculture with GCs significantly increased lipid droplets (0.50 ± 0.06/cell vs. 4.61 ± 0.98/cell, P < 0.01 vs. control, n = 90) and mitochondria with tubular cristae (3.14 ± 0.63/cell vs. 15.71 ± 1.45/cell, P < 0.01 vs. control, n = 90) in SC (Fig. 1, C and D), indicating the acquisition of active steroid synthetic capacity, a phenomenon that is well established in TCs. In contrast to SC, SM cultured alone contained abundant lipid droplets and mitochondria (Fig. 1E) and the numbers of lipid droplets (3.3 ± 0.66/cell vs. 10.0 ± 1.27/cell, P < 0.05 vs. control, n = 60) and mitochondria (13.6 ± 1.25/cell vs. 23.0 ± 1.93/cell, P < 0.05 vs. control, n = 60) were increased in the presence of GCs (Fig. 1, E and F), although to a lesser extent than for that observed in SC.


Figure 1
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  		FIG. 1. Scanning and transmission electron microscopy of cultured ovarian stromal cells. Stromal cells isolated from ovarian cortex (SC) and ovarian medulla (SM) were cultured with or without granulosa cells (GCs) for 48 h. The cultured cells (60–90 cells) were examined with a scanning electron microscope (A, B x15 000; bar = 50 µm.) and a transmission electron microscope (C, D, E, F x10 000; bar = 1 µm). SC coculturing with GCs yielded secretory products (arrow) and filopodia (arrowhead) on the cellular surfaces (B). Coculturing with GCs significantly increased lipid droplets (arrow) and mitochondria (arrowhead) in SC (D)

Androgen Production by Ovarian Stromal and Theca Cells Cultured with or Without Granulosa Cells

Isolated SC and SM were cultured alone or in the presence of GCs for up to 48 h to determine the influence of GCs on steroidogenesis by stromal cells. Androstenedione concentration in the spent media was determined. Both SC and SM cultured alone in serum-free medium produced negligible amounts of androstenedione (<0.2% of that by TCs; Fig. 2) and did not respond to hCG (0.01–1 IU/ml), whereas hCG augmented androstenedione production by TCs in a concentration-dependent manner (data not shown). Although GCs were unable to produce androstenedione (data not shown), they significantly increased androstenedione production by SC (Fig. 2A) and SM (Fig. 2B) after 24 h under coculture conditions (P < 0.05); they had no such effect on TCs (Fig. 2C).


Figure 2
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  		FIG. 2. Effect of GCs on androstenedione production by cultured ovarian stromal cells. Stromal cells isolated from the ovarian cortex (SC; A) or ovarian medulla (SM; B) and theca cells (TCs; C) were cultured without (open bar) or with (solid bar) GCs for up to 48 h, and the concentrations of androstenedione in the spent media were measured. Data are the mean ± SEM of three or four different experiments. Asterisks indicate a significant difference between single-cultured and cocultured cells (P < 0.05)

Ovarian stromal cell differentiation is characterized by increased gene expression for androgenic factors [26]. In contrast to TCs, RT-PCR analysis indicates that the levels of mRNA for CYP17A1, CYP11A1, HSD3B1, LHCGR, STAR, and NR5A1 in freshly prepared SC and SM were low and not significantly different (Fig. 3). To determine the effect of GCs on stromal cell differentiation, the levels of mRNA for CYP17A1, CYP11A1, HSD3B1, LHCGR, STAR, and NR5A1 in the single-cultured and cocultured stromal cells (SC or SM) in the serum-free medium were measured using semi-quantitative RT-PCR assays (Fig. 4). Expression of CYP17A1 and CYP11A1 mRNA but not of HSD3B1, STAR, and NR5A1 mRNA in both SC and SM increased with the duration of culture (P < 0.05 vs. control; Fig. 4, A, B, and E) and was not affected by coculture with GCs (Fig. 4E). In contrast to stromal cells, the mRNA levels of CYP17A1 and STAR in TCs decreased gradually with the duration of culture (P < 0.05 vs. control; Fig. 4, C and E). Also, GCs did not alter CYP17A1, CYP11A1, HSD3B1, LHCGR, STAR, and NR5A1 mRNA contents in TCs (Fig. 4, C and E).


Figure 3
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  		FIG. 3. Characterization of levels of mRNA in isolated ovarian stromal and theca cells. Total RNA was extracted from freshly isolated stromal cells from ovarian cortex (SC) or medulla (SM) and from theca cells (TCs). Messenger RNA expression for CYP17A1 (30 cycles), CYP11A1 (30 cycles), HSD3B1 (30 cycles), LHCGR (30 cycles), STAR (30 cycles), NR5A1 (30 cycles), VEGF (30 cycles), and ARBP (internal control; 25 cycles) were examined using RT-PCR assays. Representative images from three or four independent experiments are shown


Figure 4
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  		FIG. 4. Effect of granulosa cells (GCs) on mRNA expression in cultured ovarian stromal cells. Isolated stromal cells from ovarian cortex (SC), ovarian medulla (SM), and theca cells (TCs) were cultured with or without GCs for up to 48 h, and the levels of mRNA for CYP17A1 (stromal cells 35 cycles; TCs 30 cycles), CYP11A1 (stromal cells 35 cycles; TCs 30 cycles), HSD3B1 (stromal cells 35 cycles; TCs 30 cycles), LHCGR (stromal cells 35 cycles; TCs 30 cycles), STAR (35 cycles), NR5A1 (stromal cells 35 cycles; TCs 30 cycles), and VEGF (30 cycles) in the cultured cells were measured using RT-PCR assays. Each transcript level of target genes was normalized on the basis of the level of ARBP (25 cycles). Representative images (A-D) from three or four independent experiments and semi-quantitative data (E) of the mRNA contents (fold of control [the cells cultured alone for 24 h]) are shown. Mean ± SEM of three or four independent experiments. Bars with different letters are significantly different between culture periods, within culture type (a vs. b and x vs. y, P < 0.05). Asterisks indicate a significant difference between single-cultured and cocultured cells (P < 0.05). GCs enhanced LHCGR mRNA expression significantly in SC (A) but not in SM (B) or TCs (C). TCs did not alter the levels of mRNA for LHCGR in SC (D)

Expression of LHCGR in SC and SM was lower than that in TCs (Fig. 3). The GCs significantly enhanced LHCGR mRNA expression in SC (P < 0.05 vs. control) but not in SM after 24 h of culture (Fig. 4, A, B and E). The stimulatory effect on LHCGR mRNA expression in SC was specific to GCs because coculture with TCs did not alter expression of LHCGR mRNA in SC (Fig. 4D). In addition, this response appeared to be specific to LHCGR, because VEGF mRNA abundances were not different among the three cell types, regardless of the presence of GCs (Fig. 3 and Fig. 4E), despite known differences in the vascularization of the tissues [1] from which the cells were isolated.

Regulation of LH Responsiveness of Ovarian Stromal Cells by Granulosa Cells

To provide a more quantitative assessment of mRNA for LH receptors, real-time PCR analyses of LHCGR mRNA in stromal cells (SC and SM) and TCs were performed. Expression of mRNA for LHCGR was not detected in freshly isolated SM and SC during 35 cycles, although it was abundant in TC (0 h; Table 2). The level of mRNA for LHCGR in SC was high compared with SM, and GCs significantly enhanced LHCGR expression in SC (P < 0.05 vs. control) but not in SM or in TCs after 48 h of culture (48 h; Table 2).


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  		TABLE 2. Effect of GC on LHCGR mRNA expression in cultured ovarian stromal and theca cells.*

To determine whether the GC-induced increase in LHCGR expression in SC is associated with increases in functional LH receptors, SC were cultured alone or cocultured with GCs in serum-free medium for 24 h and were subsequently treated with hCG (0–1 IU/ml) for 30 min (cAMP assay; Fig. 5A) or 48 h (androgen assay; Fig. 5B), respectively. Concentrations of cAMP in the extracted SC cells and androstenedione in the spent media were measured. For comparison, the same experiment was performed with SM. Although no effect was apparent on single-cultured SC, hCG significantly increased cAMP and androstenedione production by SC cocultured with GCs in a concentration-dependent manner (P < 0.05; Fig. 5, A and B). In contrast, hCG had no effect on cAMP or androgen production by SM, regardless of the culture conditions.


Figure 5
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  		FIG. 5. Effect of GCs on hCG responsiveness of cultured ovarian stromal cells. To determine whether stromal cells synthesize increased amounts of cAMP and androstenedione in response to hCG stimulation when they are cocultured with GCs, isolated stromal cells from ovarian cortex (SC) and ovarian medulla (SM) were cultured with or without GCs for 24 h and then were treated with increasing concentrations of hCG (0.01–1 IU/ml) for 30 min (cAMP assay; A) or 48 h (androstenedione assay; B), respectively. The concentrations of cAMP in the extracted cells and androstenedione in the spent media were measured. Data are the mean ± SEM of three or four different experiments. Asterisks show a significant difference from the control (stromal cells cocultured with GCs in the absence of hCG stimulation; P < 0.05)

DISCUSSION

The importance of follicular cell-cell interaction during early development is poorly understood. Results of the present study demonstrate that GCs facilitate ovarian stromal cell differentiation, thereby increasing expression of mRNA for LHCGR and hCG-induced cAMP and androgen production without detectable changes in the expression of androgenic enzymes of the cell. These findings are consistent with the notion that GCs play an important role in differentiation from stromal cells to TCs during folliculogenesis.

In the present study we have demonstrated that GCs play an important role in the differentiation of SC, as evidenced by the increased number of lipid droplets and mitochondria—both of which are known to be essential for increased steroid production and enhanced basal androgen production—but no detectable change in the expression of steroidogenic enzymes. A similar increase in lipid droplets was observed when mesenchymal stem cells were forced to differentiate into androgen-producing cells (unpublished results). These findings raise the possibility that the function of GCs during early follicular development is necessary to provide an adequate substrate (e.g., steroids, growth factors, cytokines, and extracellular matrix) to maintain sufficient basal androgen biosynthesis. In the presence of GCs, stromal cells also exhibited increased expression of mRNA for LHCGR and LH responsiveness, two well-established differentiated functions of TCs in the bovine ovary [27]. In contrast, freshly isolated TCs, but not SC, expressed a high abundance of mRNA for LHCGR and CYP17A1 (another thecal biomarker). Taken together, these findings support the hypothesis that the GCs are important for functional differentiation of SC to TCs, although the nature of this cell-cell interaction remains to be determined. However, it is possible that this might be mediated through secretion and action of small molecules (<12.5 kDa; based on the membrane pore size). As the follicles transit from the preantral to the early antral stage, they acquire a highly differentiated stromal-theca cell layer containing the maximum steroidogenic capacity (high expression of key steroidogenic enzymes), and the role of GCs might be changed from a stimulatory one (i.e., gene activation) to a supportive one (i.e., provision of substrate). On the other hand, TCs lost their differentiated phenotype: CYP17A1 expression of mRNA during the culture period. The GCs stimulated neither androgen production nor mRNA expression in TCs, indicating that once stromal cells differentiate into TCs they can be maintained by LH, but not by GC, in terms of androstenedione production.

A number of studies have addressed theca-interstitial cytodifferentiation using the immature hypophysectomized rat model. Although this cell culture model is excellent for obtaining an adequate number of homogeneous theca-interstitial cells for quantitative biochemical studies, the theca-interstitial cells already express LHCGR mRNA and respond to hCG with increased androgen production [26, 28]. In contrast, the thecal precursor cells lack LHCGR mRNA, and TC recruitment occurs when the cells express genes of LH receptor and androgenic enzymes. In the present study, bovine ovarian stromal cells, which had neither LHCGR mRNA nor hCG responsiveness, were used to examine the process by which stromal cells differentiate into steroidogenic cells and acquire the capacity to respond to gonadotropins. We have demonstrated that Sc are less capable of androgen biosynthesis in vitro when compared to TCs but are nonresponsive to hCG in vitro unless cocultured with GCs. As Sc become highly differentiated, similarly to those that were cocultured with GCs, they acquire functional characteristics that resemble those of TCs and respond to hCG with a marked increase of androgen production.

It will be of interest to determine the reason for the apparent independence of CYP17A1 expression in SC of the influence of GCs if the above-mentioned contention is correct. In the present in vitro study, expression of mRNA for CYP17A1and CYP11A1 increased with the duration of culture even in single-cultured SC, suggesting the presence of an unknown factor(s) suppressing the expression of CYP17A1 and CYP11A1 mRNA in vivo, and the disappearance of this inhibitory effect might have resulted in the increased expression of CYP17A1 mRNA in the stromal cells when they were isolated and cultured alone. Previous studies suggested that the oocyte also is a source of factors that induce the expression of CYP17A1 in TCs. Whereas expression of CYP17A1 mRNA was absent in the theca compartment of growth differentiation factor-9 (GDF-9) null mice [29], administration of recombinant GDF-9 enhanced the ovarian CYP17A1 content and CYP17A1 immunostaining in TCs [30]. Taken together, these findings raise the interesting possibility that an oocyte-derived factor(s) might be involved in the upregulation of CYP17A1 mRNA during functional differentiation from SC as they develop into TCs in vivo and in overriding the suppressive effect of the unknown inhibitory factor(s).

We observed in the present study that the expression of mRNA for HSD3B1, STAR, and NR5A1 in SC was not affected by the presence of GCs. One possible explanation is that granulosal factor(s) which otherwise regulates expression of mRNA for these steroidogenic factors was not able to pass through the collagen membrane of our culture system. Using the immature hypophysectomized rat model, Huang et al. [31] reported that insulin-like growth factor I (IGF-I) increased LHCGR, CYP11A1, and HSD3B1 mRNA levels in thecal-interstitial cells. The combination of IGF-I plus kit ligand (KL) increased expression for LHCGR, STAR, CYP11A1, HSD3B1, and CYP17A1 mRNAs and androgen production in these cells [31]. Using the bovine ovarian organ culture model, Parrott and Skinner [32] reported that KL stimulated ovarian stromal cell proliferation but had no effect on androgen production, whereas estrogen increased stromal cell production of androgen. Only factors less than 12.5 kDa can pass through the collagen membrane; therefore, the possible involvement of KL (which does not pass through the membrane) could not be excluded. We speculate that GC-induced functional differentiation of SC during formation of TC is mediated in part through low-molecular weight granulosal factors that alter cellular morphology and upregulate LHCGR expression, and that the changes in steroidogenic factor(s) such as HSD3B1, STAR, and NR5A1 are dependent on larger factor(s) secreted from GCs. Further studies are necessary to determine the identity of the factor(s) and the mechanism of influence upon stromal cell differentiation into TCs.

In the present study we have shown that, in contrast to SC, SM contained abundant lipid droplets and mitochondria and are less influenced by the presence of GCs (e.g., LHCGR mRNA expression), indicating that SM already possess steroidogenic competence but not the capability of differentiating into TCs in response to GC-derived factor(s). All stromal cells are assumed to arise from a population of unspecialized mesenchymal stem cells [33]. Consistent with this notion is the observation that the developing fetal ovine ovary is steroidogenically active from Day 30 to Day 75 [34–36], with steroidogenically active cells being abundant from Days 35–45 but localized to the outer medulla/inner cortex region. At Day 55, the steroidogenically active cells are restricted entirely within the stromal cells of the ovarian medulla, particularly in cells of the mesonephric-derived cell streams.

In summary, using a coculture system we have examined the possibility that GC-stromal cell interaction plays a role in the differentiation of ovarian stromal cells into TCs. We have demonstrated that GCs from small antral follicles induce the functional differentiation and the acquisition of LH responsiveness in cortical stromal cells, but not in medullary stromal cells. The TCs first appear in bovine follicles at the secondary stage, and the TC layer forms during the preantral to early antral stage of ovarian follicular growth [7, 11]. Results of the present study suggest that GCs of secondary follicles might have the same type of interaction with cortical stromal cells that surround them. Whether the continued growth of the TC layer is caused by continued effects of GCs on additional cortical stromal cells and/or by simple proliferation of existing TCs remains to be determined. Nevertheless, our findings are consistent with the notion that GC-SC interaction plays a role in TC recruitment during early follicular development.

ACKNOWLEDGMENTS

We would like to thank Dr. Abraham Amsterdam and Carmen Cheung for reviewing the manuscript and for offering their helpful comments, and Mikiko Misawa for excellent technical assistance and earnest cooperation. We are grateful to National Hormone & Peptide Program (Harbor-UCLA Medical Center; Torrance, CA) and Serono (Geneva, Switzerland) for providing hCG and recombinant human LH (LHadi), respectively. We also thank the Kanazawa Meat Inspection Office (Kanazawa, Japan) for allowing collection of the bovine ovaries used in these experiments.

FOOTNOTES

2 Correspondence: FAX: 81 776 61 8117; kotsujif{at}fmsrsa.fukui-med.ac.jp Back

1 Supported by the Ministry of Education, Culture, Sports, Science, and Technology, Japan and the Canadian Institutes of Health Research. Back

Received: 19 December 2005.

First decision: 26 January 2006.

Accepted: 14 August 2006.

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