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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
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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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 (
5ß1,
iibß3, and most all5ß
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.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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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 = 
(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: 
(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. 3
. 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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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 
2 test and linear regression analysis.
Regardless of the statistical test, only P < 0.05 was
considered significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). 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 1). This large increase in the surface
area of the ovary is dramatically illustrated by scanning electron
microscopy (Fig. 2).
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). 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 1).
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 1, 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. 4). 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. 4).
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). 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. 6). Similarly, the functional LH receptor
mRNA was not found in RNA samples prepared from ROSE-179 cells (Fig. 6). As expected, functional LH receptor mRNA was detected within RNA
samples derived from whole immature rat ovaries (Fig. 6). 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. 6).
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). 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. 7b).
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and 9a).
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. 9b). The separating cells continued to move apart over the culture
period, reaching an average distance between epicenters of 9.5 µm
(Fig. 9b). 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. 8 and 9a). This
transient reduction in cell contact was associated with the cells
undergoing cell division (Fig. 8). In the absence of HGF
(i.e. control conditions), 97 ± 2% of all cell
contacts were maintained regardless of substrate (Fig. 9a). 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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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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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.
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Acknowledgments |
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Footnotes |
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2 Supported by fellowships from Krebsliga des Kantons (Zurich,
Switzerland) and Deutsche Forschungsgemeinschaft (HE 288212–1).
Received October 28, 1998.
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