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Endocrinology Vol. 140, No. 6 2908-2916
Copyright © 1999 by The Endocrine Society

ARTICLES

Hepatocyte Growth Factor Induces Rat Ovarian Surface Epithelial Cell Mitosis or Apoptosis Depending on the Presence or Absence of an Extracellular Matrix1

Stefanie Hess2, Rita Gulati and John J. Peluso

Department of Obstetrics and Gynecology, University of Connecticut Health Center, Farmington, Connecticut 060030

Address all correspondence and requests for reprints to: John J. Peluso, Ph.D., Department of Obstetrics and Gynecology, University of Connecticut Health Center, Farmington, Connecticut 060030. E-mail: peluso{at}nso2.uchc.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The present studies showed that sequential treatment with equine CG (eCG) and hCG not only induced an increase in ovarian weight, but also caused an estimated 4.6-fold increase in the number of ovarian surface epithelial cells. In addition, eCG-hCG treatment increased ovarian hepatocyte growth factor (HGF) messenger RNA levels. These studies also demonstrated that rat primary ovarian surface epithelial cells as well as a cell line derived from rat ovarian surface epithelium (i.e. ROSE-179 cells) do not express the LH (hCG) receptor. Both of these cells express c-Met, the receptor for HGF. To assess the effects of hCG and HGF on ovarian surface epithelial cell mitosis, ROSE-179 cells were cultured for 24 h in serum-supplemented medium on either glass or the synthetic fibronectin-like extracellular matrix protein, pronectin (RGD). The cells were then cultured for 24 h in serum-free medium in the presence or absence of hCG or HGF. The numbers of cells at 2, 24, and 48 h of culture were determined. The percentage of apoptotic cells was assessed by in situ DNA staining at 48 h of culture. In the serum-supplemented medium in the presence or absence of RGD, the number of ROSE-179 cells doubled. In serum-free medium, cell proliferation was reduced, and the percentage of apoptotic nuclei ranged between 10–15% regardless of the substrate. Neither mitosis nor apoptosis was influenced by hCG in the presence or absence of RGD. For ROSE-179 cells cultured in serum-free medium on RGD, HGF induced mitosis, resulting in a 2.8 ± 0.2-fold increase in cell number compared with the 24 h control values. On a glass substrate in serum-free medium, HGF did not induce mitosis, but increased the percentage of apoptotic nuclei. Time-lapse photographic analysis revealed that on RGD, cells undergoing HGF-induced mitosis showed a transient reduction in cell contact. On glass, HGF caused many cells to completely lose contact and separate from each other. Collectively, these data suggest that in vivo gonadotropins stimulate HGF expression and ovarian surface epithelial cell proliferation. Based on in vitro studies, it is likely that the mitogenic action of hCG is mediated by HGF. However, HGF only induces mitosis in the presence of an extracellular matrix.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE RAT OVARY is covered by a single layer of epithelial cells (1). This continuous epithelium layer is maintained during periods of rapid follicular growth and corpus luteum formation. As both follicular growth and luteal formation are under the control of gonadotropins (2), it has been assumed that gonadotropins stimulate the ovarian surface epithelial cells to proliferate. However, it is not clear that gonadotropins act directly on ovarian surface epithelial cells to stimulate mitosis.

Another mechanism to account for the ability of gonadotropins to promote ovarian surface epithelial cell mitosis could involve the synthesis and secretion of various growth factors. Numerous studies have shown that gonadotropins act on both granulosa and thecal cells to stimulate the synthesis of several growth factors (3). Recently, hCG (4) as well as estrogen (5) have been shown to promote the expression of hepatocyte growth factor (HGF) within the ovary. In non-ovarian cells, HGF regulates many physiological responses, such as mitosis, motogenesis (i.e. cell movement), morphogenesis, and apoptosis (6, 7, 8). Which response is elicited may depend on the presence of other hormones and growth factors. In addition, cell-extracellular matrix and cell-cell interactions could influence HGF’s biological action.

Interestingly, the basement membranes within mammalian ovaries are composed of laminin, type IV collagen, and fibronectin (9). All of these proteins contain the Arg-Gly-Asp (RGD) tripeptide sequence that is recognized by several integrins ({propto}5ß1, {propto}iibß3, and most all{propto}5ß integrins) (10, 11). Once the integrins, which are expressed at the cell surface, bind to the RGD sequence, a signal transduction cascade is stimulated that modulates many cellular processes, including cell viability (12). In this light, it is interesting that at the time of hCG-induced ovulation, some ovarian surface epithelial cells undergo mitosis, whereas others undergo apoptosis (13, 14). The mechanism that accounts for ovarian surface epithelial cells undergoing these very different physiological processes in response to the exact same hormone stimuli remains to be determined, but may be related to the presence or absence of a basement membrane. Therefore, a series of studies was designed 1) to estimate the rate at which ovarian surface epithelial cells proliferate in response to gonadotropin treatment in vivo, 2) to characterize the pattern of LH receptor and c-Met expression using RT-PCR, 3) to assess the mitogenic action of hCG and HGF using a cell line derived from rat ovarian surface epithelium, and 4) to determine the effect of an extracellular matrix on the mitogenic, motogenic, and apoptotic actions of HGF in vitro.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
In vivo studies
Animals. Immature female Wistar rats (22 days of age) were obtained from Charles River Laboratories, Inc. (Wilmington, MA), and housed under controlled conditions of temperature, humidity, and photoperiod (12 h of light, 12 h of darkness; lights on at 0700 h). At 23 days of age, rats were anesthetized in CO2 and cervically dislocated between 0930–1000 h. Other rats were injected ip with 20 IU equine CG (eCG; Sigma Chemical Co., St. Louis, MO), and their ovaries were removed 48 h later. A third group was pretreated with eCG, then injected ip with 10 IU hCG (Sigma Chemical Co.) 48 h later. Their ovaries were removed 72 h after hCG. In this model, ovulation occurs between 12–14 h after hCG. Control rats were also autopsied at 28 days of age. This protocol was approved by the animal care committee of the University of Connecticut Health Center.

Estimation of ovarian surface epithelial cell proliferation in vivo. To assess ovarian surface epithelial cell proliferation in vivo, the surface area of the entire ovary was estimated before and after gonadotropin treatment. Next, the average surface area of an ovarian surface epithelial cell was determined. By dividing the surface area of the ovary by the average surface area of an ovarian surface epithelial cell, the number of ovarian surface epithelial cells per ovary was estimated. The procedures used to monitor these two end points are outlined below.

Ovarian surface area estimates. The ovaries were fixed in 10% buffered formalin and embedded in paraffin. Serial 10-µm sections were cut through the entire ovary. These sections were mounted on poly-L-lysine-coated slides and stained with hematoxylin (Diff-Quick Stain Set, Baxter, Miami, FL). An image of every fifth section was captured using a Dage 72 CCD camera (Michigan City, IN) and was stored in a computer. Using IP Lab Spectrum software (Signal Analytics Corp., Vienna VA), the perimeter of every fifth ovarian section was measured and used to derive an average radius. The surface area of the entire ovary was calculated using the following formula for the curved surface of a cone: ovarian surface area = {Sigma} (curved surface area between every fifth section) + area of the first section + area of the last section. The curved surface between every fifth section was calculated using the following formula: {pi}(r1 + r2) x [rad]h2 + (r1 - r2)2, where r1 is the radius of the top section, r2 is the radius of the bottom section, and h is the distance between sections.

Ovarian surface epithelial cell area measurements. Ovaries were prepared for examination under a scanning electron microscope using standard protocols (15). Low magnification images were taken of the entire ovary, whereas higher magnification images were taken from three different areas of each ovary. All images were stored in a computer. The surface area of epithelial cells that covered the surface in three randomly selected areas within each ovary was determined using IP Lab Spectrum software (Signal Analytics Corp.). The surface area was determined from high magnification micrographs, as shown in Fig. 3Go. The percentage of small compacted and large flattened cells in each micrograph was estimated. Then the surface area of between 50–100 cells of each type was measured. The average surface area of an ovarian surface epithelial cell was calculated according to the ratio of small and large cells. In total, ovaries from three different rats were analyzed for each treatment group.



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  		Figure 3. A higher magnification of the surface epithelium of immature rat ovary. Note that there are various sized ovarian surface epithelial cells. The largest cells are associated with the apex of the follicles, and smaller cells are found along the lateral surfaces.

 
Isolation of primary ovarian surface epithelial cells. Rat ovarian surface epithelial cells were isolated from 25-day-old Wistar rats as described by Quirk et al. (16) with slight modifications. Two or three rats were used for each preparation as outlined. First, the ovary was incubated with a dispase/collagenase mixture [2.4 U/ml dispase (Boehringer Mannheim, Mannheim, Germany) and 0.5% collagenase (Sigma Chemical Co.)] for 35 min at 37 C. The ovaries were removed and washed twice with DMEM-Ham’s F-12. The remaining cell suspension was centrifuged at 300 x g for 5 min, and the cell pellet was washed twice with DMEM-Ham’s F-12 medium before RNA extraction. At least three preparations were assessed in these studies. To assess the purity of the cell preparation, 200 µl of the cell suspension were centrifuged in a Shandon cytocentrifuge at 1100 rpm for 5 min, fixed in 10% formalin, and stained for cytokeratin using a monoclonal antipan cytokeratin (Sigma Chemical Co.) and the diaminobenzidine substrate kit (Vector Laboratories, Inc., Burlingame, CA).

Detection of HGF, LH receptor, and c-Met messenger RNA (mRNA) by RT-PCR. RNA was isolated from fresh ovarian tissue, isolated ovarian surface epithelial cells, and ROSE-179 cells with the Ultraspec RNA Isolation System following the manufacturer’s protocol (Biotex Laboratories, Inc., Houston, TX). RT was performed using 500 ng random primers (Life Technologies, Gaithersburg, MD)/4 µg RNA (10 µg RNA for experiments with treated ovaries) in a total volume of 12 µl. The mixture was incubated at 70 C for 10 min and then quickly chilled on ice. Seven microliters of a mastermix containing 2 µl PCR buffer (10-fold concentrated; Sigma Chemical Co.), 1 µl MgCl2 (50 mM; Life Technologies), 1 µl deoxy-NTP mix (10 mM each; Life Technologies), 2 µl dithiothreitol (0.1 M; Life Technologies), and 1 µl diethylpyrocarbonate-treated H2O were added and incubated for 25 min at 25 C. Two hundred units reverse transcriptase (Superscript, Life Technologies) per reaction were added, and the reaction was incubated at 25 C for 10 min and then at 42 C for 50 min. The reverse transcriptase was inactivated at 70 C for 15 min, and the solution was cooled on ice for 2 min.

Two microliters of the RT reaction were used for PCR amplification. PCR for detection of HGF in treated ovaries was performed in a volume of 100 µl containing 1.5 mM MgCl2 and 2.5 U Taq polymerase (Sigma Chemical Co.) for 35 cycles (initial denaturation at 94 C for 4 min, denaturation at 94 C for 1 min, annealing at 52 C for 30 sec, elongation at 72 C for 1 min, and final elongation for 10 min). For detecting LH receptor, c-Met, and hypoxanthine phosphoribosyl transferase (HPRT), the following hot start procedure was performed to avoid primer dependent PCR artifacts. Taq polymerase (2.5 U in a total volume of 100 µl) was added after denaturing the reactions at 94 C for 3 min. PCR was performed for 30 or 45 cycles depending on the experiment. Each cycle consisted of denaturing at 94 C for 1 min, annealing at 52 C for 90 sec, elongation at 72 C for 90 sec, and a final elongation step for 10 min. Oligonucleotide primers were designed to anneal at different exons to distinguish template source, complementary DNA (cDNA), and contamination with chromosomal DNA, respectively. The following primers were selected from homologous regions of the human and rat HGF cDNA sequence (17): antisense, 5'-CATCAAAGCCCTTGTCGGGATA-3' (nucleotides 792–813); and sense, 5'-ACAGCTTTTTGCCTTCGAGCTA-3' (nucleotides 523–544). For the LH receptor the following primers were used (18, 19): antisense, 5'-TAACTGTGCTTTCGCATTG-3' (nucleotides 962–980); sense, 5'-CGAGTCCCAGCTCTGAGACA-3'(nucleotides 53–72). The primers for c-met were: antisense, 5'-TGGCTCCCAGGGCACTCACTACAC-3' (nucleotides 527–550); sense, 5'-GGCCCGTGGTGGAACACCCAGATT-3' (nucleotides 260–283) (20). In parallel, PCR of the HPRT was performed using the same amount of cDNA from each sample as a template. Primers were designed according to a previous report (21), with the exception of a 1-bp exchange for the rat HPRT-coding sequence, as follows: 5'-GTCAAGGGCATATCCAACAACAAAC-3' (nucleotides 656–680; antisense) and 5'-CCTGCTGGATTACATTAAAGCGCTG-3' (nucleotides 329–353; sense) (22). An aliquot of the PCR was separated on a 1.5% agarose gel containing ethidium bromide (0.5 µg/ml) and visualized under UV light. The RT-PCR reaction resulted in a HGF fragment of 291 bp, functional LH receptor was detected by the presence of a 928 bp fragment, the c-Met RT-PCR reaction yielded a 291-bp fragment, and the HPRT fragment was 352 bp. This protocol was repeated on at least two separate sets of RNA samples isolated from each treatment.

In vitro studies
Rat ovarian surface epithelial (ROSE-179) cell culture. ROSE-179 cells were generously provided by Dr. Robert Burghardt of Texas A&M University (College Station, TX) and cultured in DMEM-Ham’s F-12 medium (Sigma Chemical Co.), supplemented with 5% FBS (HyClone Laboratories, Inc., Logan, UT) (23). For studies involving proliferation or apoptosis, ROSE-179 cells were plated in eight-chamber glass Lab-Tek slides (Nunc, Inc., Naperville, IL) at 30,000 cells/400 µl. The Lab-Tek wells were coated with the RGD substrate by diluting pronectin (Sigma Chemical Co.) 1:10 with sterile PBS and adding enough of this solution to cover the entire culture surface of each well. After a 2-h incubation, the Lab-Tek slides were washed twice with PBS before plating with the ROSE-179 cells in serum-supplemented medium. Recombinant human HGF (Genentech, Inc., South San Francisco, CA) was used at a concentration of 4 ng/400 µl, and hCG (Sigma Chemical Co.) was used at a concentration of 10 IU/400 µl.

Assessment of mitosis and apoptosis. ROSE-179 cell proliferation was determined using an in situ cell counting procedure (24). Briefly, cells were cultured in serum-supplemented medium. After 2 h, the number of cells in four different large grids within each Lab-Tek well was counted. After 24 h, the cells were washed with serum-free medium, and the number of cells in the same large grids within each Lab-Tek well was counted. The cells were then incubated for 24 h in either serum-free medium supplemented with hCG or HGF with or without RGD matrix. After this 24-h incubation, the cells were counted again. Cell proliferation was expressed as a fold increase in cell number (i.e. 24-h count/2-h count and 48-h count/24-h count). The cells were then stained with hydroethidine and observed under fluorescent optics to assess apoptosis, as previously described (24). A cell was considered to be apoptotic if its nucleus had fragmented, forming apoptotic bodies (24).

Monitoring ROSE-179 cells using time-lapse photography. Time-lapse studies were conducted as previously described (24). ROSE-179 cells were cultured in serum-supplemented medium for 24–48 h, then washed with serum-free medium and cultured in serum-free medium with or without HGF in the presence or absence of RGD matrix. Sequential images were collected at 30-min intervals over a 4-h period. To ensure an accurate assessment of cell contact, only aggregates of three cells or fewer were examined. A cell contact was considered lost when the cells were completely detached. The number of initial cell contacts that remained at each time interval was counted, and a percentage was calculated. Experiments were performed on 4 different days, and at least 50 cell contacts were examined.

The images from the time-lapse studies were also analyzed to assess the motogenic action of HGF (24). For this analysis, each ROSE-179 cell that became completely detached from an aggregate of cells was studied. The distance between epicenters of the cells that detached was measured at 30-min intervals over a 4-h period using IP Lab Spectrum software (Signal Analytics Corp.). All time intervals were normalized to the time at which cell attachment was lost. As not all cells lost attachment at the same time, not all detached cells were studied for a full 4-h period. However, to be included in the analysis, cells were observed for at least 2 h.

Statistical analysis
Each experiment involving proliferation or apoptosis was conducted in duplicate and replicated three times. The data from these experiments were analyzed by ANOVA after determining that the values were normally distributed. The data were subsequently analyzed by a Student-Newman-Keuls multiple range test. Changes in cell contact were analyzed by {chi}2 test and linear regression analysis. Regardless of the statistical test, only P < 0.05 was considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effect of gonadotropins on ovarian weight, ovarian surface area, and number of ovarian surface epithelial cells
Ovarian weight of immature rats remained relatively constant between 23 and 28 days of age. Sequential treatment with eCG and hCG resulted in 4- and 12-fold increases in ovarian weight after 2 and 5 days of gonadotropin treatment, respectively (Fig. 1Go). Similarly, morphometric studies revealed that eCG-hCG treatment increased ovarian surface area by 3.5- and 10-fold after 2 and 5 days, respectively (Table 1Go). This large increase in the surface area of the ovary is dramatically illustrated by scanning electron microscopy (Fig. 2Go).



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  		Figure 1. The effects of eCG and hCG on ovarian weight. Values in this and subsequent graphs are expressed as the mean ± SE. Note that at some time points the SE was too small to be observed on the graph.

 

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  		Table 1. The effect of sequential eCG and hCG treatment on ovarian surface area, ovarian surface epithelial cell surface area, and cell number

 


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  		Figure 2. The effects of eCG and hCG on ovarian morphology as observed by scanning electron microscopy. The upper panel shows an immature ovary; the middle panel shows an eCG-primed ovary, and the lower panel shows an ovary after eCG and hCG treatment. All figures are shown at the same magnification.

 
Scanning electron microscopic analysis further showed that the ovarian surface was covered by various sized epithelial cells. The largest epithelial cells were very flat and were observed on top of the follicles and corpora lutea, whereas smaller compact cells were located within the crevices between the follicular and/or luteal structures (Fig. 3Go). The percentages of small compact and large flattened cells were estimated from the micrographs. The percentage of small compact cells that covered the surface of the nongonadotropin-treated ovary was approximately 50%. After gonadotropin treatment, this percentage decreased to about 30%. In immature rats the average epithelial cell area was 92 ± 2 µm2. After eCG treatment, the average epithelial cell area increased to 194 ± 8 µm2 (P < 0.05). The average area of the epithelial cells did not change in response to subsequent treatment with hCG (Table 1Go).

Using the estimates of ovarian surface area and average ovarian surface epithelial cell area, the relative number of ovarian surface epithelial cells present after gonadotropin treatment was calculated. As shown in Table 1Go, the number of ovarian surface epithelial cells increased 1.6- and 4.6-fold within 2 and 5 days of gonadotropin treatment.

Expression of HGF in rat ovaries
HGF mRNA was detected in total ovarian RNA samples from all treatment groups (Fig. 4Go). The levels of HGF mRNA were barely detectable in control ovaries, but were readily detected in ovaries from rats treated with gonadotropins. HPRT mRNA was detected in all samples (Fig. 4Go).



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  		Figure 4. The detection of HGF mRNA (upper panel) and HPRT mRNA (lower panel) by RT-PCR within immature ovaries treated with eCG or eCG plus hCG. The positive control represents RNA isolated from liver. For the negative control in this figure as well as Fig. 6Go, Millipore water was used to replace the DNA sample. Data shown in this figure were generated using 30 cycles of PCR.

 
Expression of LH receptor and C-Met by primary ovarian surface epithelial and ROSE-179 cells
The dispase/collagenase yielded a uniform population of cells that possessed the morphological characteristics of epithelial cells. Further, nearly 100% of these cells stained positively for cytokeratin, a marker for epithelial cells (Fig. 5Go). RT-PCR analysis of RNA isolated from this ovarian surface epithelial cell preparation failed to detect the presence of mRNA that encodes for the functional LH receptor (Fig. 6Go). Similarly, the functional LH receptor mRNA was not found in RNA samples prepared from ROSE-179 cells (Fig. 6Go). As expected, functional LH receptor mRNA was detected within RNA samples derived from whole immature rat ovaries (Fig. 6Go). In contrast, PCR fragments associated with c-Met and HPRT were detected in both of these cell preparations as well as in whole ovary extracts (Fig. 6Go).



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  		Figure 5. Primary ovarian surface epithelial cells isolated after dispase/collagenase digestion. Cells were stained in the absence (left panel) or presence of an antibody to cytokeratin, an epithelial cell marker. Nearly 100% of the cells isolated by the dispase/collagenase protocol stained for cytokeratin, indicating that they were surface epithelial cells. Cells are shown at a final magnification of x1000.

 


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  		Figure 6. The detection of LH receptor (upper panel), c-Met (middle panel), and HPRT (lower panel) mRNAs within primary ovarian surface epithelial cells, ROSE-179 cells, and immature rat ovaries. Data shown in this figure were generated using 45 cycles of PCR.

 
Effects of hCG and HGF on ROSE-179 cell proliferation and apoptosis
As would be predicted from the RT-PCR analysis, hCG did not affect the rate at which ROSE-179 cells underwent either mitosis or apoptosis regardless of whether the cells were plated on glass or RGD substrate (data not shown). In contrast to hCG, HGF treatment resulted in a 2.8-fold increase in the cell number for cells plated on RGD-coated slides. This increase in the cell number was significantly greater than that observed for all other treatment groups (P < 0.05; Fig. 7aGo). The HGF-induced increase in cell number was also greater than that observed for cells cultured in serum-supplemented medium (2.1 ± 0.1-fold increase/24 h; P < 0.05; data not shown). If cells were plated on glass, HGF increased the percentage of apoptotic nuclei compared with values in all other treatment groups (P < 0.05; Fig. 7bGo).



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  		Figure 7. The effects of the extracellular matrix, RGD peptide, and HGF on ROSE-179 cell proliferation (A) and apoptosis (B) in vitro. ROSE-179 cells were cultured on glass or RGD substrate. Proliferation was expressed as the fold increase in cell number over the 24-h period between 24–48 h of culture. *, The value is greater than those in all other groups (P < 0.05).

 
HGF regulation of cell contact and movement
Time-lapse studies showed that HGF caused about 20% of the cells that were plated on glass substrate to separate within 4 h (Figs. 8Go and 9aGo). This loss of cell contact was due to HGF stimulating cells to move apart, with the distance between the epicenter of the cells increasing an average of about 6 µm within 30 min of the cells separating (Fig. 9bGo). The separating cells continued to move apart over the culture period, reaching an average distance between epicenters of 9.5 µm (Fig. 9bGo). Conversely, cells plated on the RGD substrate and treated with HGF showed a transient reduction in the amount of surface membrane that remained in contact with other cells, but cell contact was never completely lost during the 4-h culture period (Figs. 8Go and 9aGo). This transient reduction in cell contact was associated with the cells undergoing cell division (Fig. 8Go). In the absence of HGF (i.e. control conditions), 97 ± 2% of all cell contacts were maintained regardless of substrate (Fig. 9aGo). To ensure accurate assessment of cell contact, the above data were generated by monitoring aggregates of three cells or less. However, cell within larger aggregates showed a similar response to HGF.



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  		Figure 8. Sequential changes in HGF-induced cell contact as observed by time-lapse photography. ROSE-179 cells were cultured on either glass or RGD substrates. Note that on glass in the presence of HGF, the initial cell contacts, identified by arrows in the upper left panel, were lost over the culture period. In contrast, cells plated on RGD substrate do not completely lose contact in response to HGF. Rather, HGF promotes cell division. The time after HGF exposure is shown in the lower right corner of each photograph.

 


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  		Figure 9. The effects of HGF and extracellular matrix on ROSE-179 cell contact (A) and motogenic response (B). In this study cells were cultured on either glass or RGD with serum for 24 h. Then serum was removed, and the cells were cultured in the presence or absence of HGF. In B, the change in the distance between epicenters of cells that have separated is shown on the y-axis and is plotted in relationship to the time when the loss of cell contact was initially observed. Controls were cultured on either glass or RGD substrate and were pooled, as under control conditions cell contact was maintained on both substrates.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
A continuous epithelial cell layer is maintained over the ovarian surface throughout the course of gonadotropin-induced follicular growth and luteal formation. The present study demonstrates that two physiological mechanisms are involved in this process. The first mechanism involves the regulation of epithelial cell surface area. In several mammalian species, the ovarian epithelium is composed of different sized cells (see Ref. 15 for citations). These different cell sizes appear to represent not different cell types but, rather, different phenotypes (25). Weakley (25) proposed that as the ovarian surface area increases, the epithelial cells increase in surface area as a result of the mechanical forces associated with the cells maintaining contact with each other and the basement membrane. This concept is supported by observations from several groups showing that the surface epithelial cells become flattened and distended at the apex of the ovulatory follicle (see Ref. 15 for citations). A similar response to mechanical forces could account for the increase in the average surface area of the ovarian surface epithelial cells during eCG-induced follicular growth. However, the present data indicate that an increase in cell surface area is not the only mechanism required to maintain a continuous ovarian surface epithelium during follicular development. In fact, the increase in epithelial cell surface area only accounts for about half of the increased ovarian surface area associated with eCG-induced follicular development, with an increase in cell number accounting for the remainder. In addition, all of the hCG-induced increase in the ovarian surface epithelium is due to an increase in cell number and not to an increase in the surface area of individual ovarian surface epithelial cells.

To maintain an intact epithelium, the epithelial cells proliferate, undergoing a nearly 5-fold increase in cell number after 5 days of gonadotropin treatment. The mechanism by which gonadotropins promote ovarian epithelial cell proliferation is unknown. From the RT-PCR studies, it appears that neither primary ovarian surface epithelial cells nor ROSE-179 cells express the mature, functional form of the LH receptor. The observation that hCG does not stimulate ROSE-179 cell mitosis is consistent with the RT-PCR analysis. Based on these data, it is likely that hCG does not act directly on rat ovarian surface epithelial cells to promote their mitosis but, rather, acts indirectly by stimulating the synthesis of ovarian growth factors such as HGF. However, hCG has been shown to stimulate mitosis of primary rabbit ovarian surface epithelial cells (26). There are at least two possible explanations for this discrepancy. First, there may be species differences between rat and rabbit ovarian surface epithelial cells. Second, the preparation of primary rabbit ovarian surface epithelial cells may have been contaminated with stromal/thecal cells, which could mediate a hCG-HGF paracrine mechanism, as proposed in the subsequent discussion.

Previous studies have shown that both hCG (4) and estrogen (5) increase ovarian HGF mRNA levels, with the thecal/stromal cells being the site of HGF synthesis (27). The present study also demonstrates that c-Met is expressed in both primary ovarian surface epithelial cells and ROSE-179 cells. Although there are numerous growth factor within the ovary that could promote ovarian surface epithelial cell mitosis, these studies show that HGF is a potent ovarian epithelial cell mitogen, inducing a nearly 3-fold increase in the number of ROSE-179 cells within 24 h. These observations are consistent with the hypothesis that in vivo HGF mediates the mitogenic action of hCG on rat ovarian surface epithelial cells by activating the c-Met receptor.

The observed mitogenic effect of HGF on ROSE-179 cells expands the paracrine role proposed by Parrott and associates (4, 27) for HGF in regulating ovarian function. To support their paracrine model, these investigators have shown that 1) HGF is expressed by the stromal/thecal cells of bovine follicles, 2) hCG stimulates HGF expression by the bovine stromal/thecal cells, and 3) HGF acts on bovine granulosa cells to promote their proliferation. In addition, HGF suppresses FSH-induced aromatase activity (4) and LH-induced thecal androgen synthesis (28). As androgen can induce follicular atresia, this action could maintain the viability of the follicles. Based on these observations, it has been proposed that during follicular development a paracrine axis exists in which gonadotropins and estrogen increase the HGF concentration within developing follicles. This increase in HGF may be important not only in stimulating granulosa cell proliferation within ovarian follicles but also in stimulating the proliferation of the ovarian surface epithelial cells that overlie them. A similar paracrine axis may be associated with the developing corpus luteum.

The mitogenic effect of HGF is only observed when ovarian surface epithelial cells are plated on a fibronectin-like extracellular matrix (RGD peptide). HGF has been shown to decrease cadherin levels (29) and increase the tyrosine phosphorylation of ß-catenin (30). These combined actions result in the loss of cell contact (31). It is important to appreciate that N-cadherin-mediated cell contact regulates a signal transduction pathway that maintains ROSE-179 cell viability (32, 33). In other systems, fibronectin-integrin binding also promotes cell survival (12). Thus, the N-cadherin- and integrin-mediated pathways may activate redundant cell survival mechanisms. This could allow N-cadherin-mediated cell contact to be disrupted during HGF-induced mitosis without resulting in cell death. This may explain why HGF stimulates mitosis when cells are on RGD substrate and apoptosis in the absence of RGD.

These data suggest that RGD binds to integrins that are apparently present within the cell membranes of ROSE-179 cells. This interaction sets in place a signal transduction pathway that converts HGF’s apoptotic and motogenic actions into a mitogenic action. The nature of this putative RGD-induced signal transduction pathway is unknown. Numerous studies have suggested that fibronectin (RGD) interacts with integrins to activate several tyrosine kinases and mitogen-activated protein kinase signal transduction pathways (34). These pathways often play essential roles in the cell survival and mitotic signal transduction cascades (35, 36). It is proposed that in ROSE-179 cells, HGF does not activate all the components of the mitotic signal transduction cascade. Rather, HGF probably induces a subset of components, including those that reduce N-cadherin-mediated cell contact. The remaining components of the mitogenic cascade are stimulated by an RGD-integrin interaction. It appears that for the ROSE-179 cell to undergo mitosis in response to HGF, both the HGF and RGD-integrin pathways must be activated. If the RGD component is absent, then mitosis is interrupted, and the ROSE-179 cells undergo apoptosis. A similar relationship between mitosis and apoptosis has been observed in several other systems (37, 38, 39). However, further studies are needed to determine whether other extracellular matrix proteins can modulate the effect of HGF on ROSE-179 cell function.

Finally, the present studies demonstrate that in the absence of extracellular matrix, HGF decreases cell contact, promotes cell migration, and induces apoptosis. This in vitro situation mimics that associated with ovulation. During the final stages of the ovulatory process, the extracellular matrix at the apex of the ovulatory follicle is degraded (40), and those ovarian surface epithelial cells at the apex undergo apoptosis (13, 14, 34). This results in a stigma through which the oocyte is released (13, 14, 34). At the same time, ovarian surface epithelial cells adjacent to the apex undergo mitosis and eventually fill in the stigma (13, 14, 34). Thus, the presence or absence of an extracellular matrix may explain why during ovulation some ovarian surface epithelial cells undergo mitosis while other undergo apoptosis in response to the exact same hormonal and growth factor milieu.


   Acknowledgments
 
The authors are indebted to Ms. Anna Papplardo for her excellent technical assistance. The authors also thank Dr. Bruce White for his thoughtful comments and suggestions, and Dr. Ralph Schwall of Genentech, Inc., for providing the human hepatocyte growth factor.


   Footnotes
 
1 This work was supported in part by NIH Grant 1-RO-1-HD-33467–01A. Back

2 Supported by fellowships from Krebsliga des Kantons (Zurich, Switzerland) and Deutsche Forschungsgemeinschaft (HE 288212–1). Back

Received October 28, 1998.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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