Infertility Clinic, Department of Gynaecology and Obstetrics, Geneva University Hospital, 1211 Geneva 14, Switzerland and 2Fondation pour Recherches Médicales, 64, avenue de la Roseraie, University of Geneva, 1205 Geneva, Switzerland
Received on June 27, 2000; revised on October 3, 2000; accepted on October 4, 2000.
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Abstract |
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Key words: Anticoagulant heparan sulfate/antithrombin III/heparan sulfate proteoglycan/ovarian granulosa cells/coagulation
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Introduction |
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Anticoagulant HSPGs (aHSPGs) are synthesized by endothelial cells and endow vessel walls with antithrombotic properties (de Agostini et al., 1990; Rosenberg and Aird, 1999
). In addition, a heparin-like activity has been detected in ovarian granulosa cells and follicular fluid (Andrade-Gordon et al., 1992
), and we have demonstrated that cultured rat granulosa cells synthesize significant amounts of aHSPGs, similar to that produced by endothelial cells (Hosseini et al., 1996
). Granulosa cell HSPG core proteins have been originally described by Yanagishita and Hascall, who reported the synthesis of a membrane-spanning and a glycosyl-phosphatidylinositol (GPI)anchored HSPGs (Yanagishita and Hascall, 1984
, Yanagishita and McQuillan, 1989
). Recently, expression of the four members of the syndecan family of HSPGs was reported in the mouse ovary, syndecan-4 being up-regulated in degenerating atretic follicles (Ishiguro et al., 1999
). Moreover, human follicular fluid was shown to contain a composite form of HSPG core protein immunologically related to perlecan (Eriksen et al., 1999
).
The inner ovarian follicle is formed by granulosa cells surrounding the oocyte and remains avascular until ovulation (Fortune, 1994). The production of aHSPGs requires the assembly of a complex biosynthetic pathway able to generate AT-binding pentasaccharide sequences (Rosenberg et al., 1997
). This synthesis in ovarian granulosa cells suggests a previously unsuspected function of aHSPGs outside from the vascular bed. At ovulation, plasma proteins leak from permeabilized vessels surrounding ovulatory follicles and fibrin gets deposited in the outer layers of the follicle (Dvorak et al., 1999
).
Granulosa cell aHSPGs could be critically expressed during the development of ovarian follicles, to prevent clotting of follicular fluid and thus contribute to the maintenance of fluidity in the environment of the oocyte. To test this hypothesis, this study evaluated the expression pattern of aHSPGs in granulosa cells in response to hormonal stimulations in vitro and in vivo. We measured the aHSPGs produced by granulosa cells following gonadotropin stimulations in vitro and simultaneously visualized aHSPGs on the cells by microscopic autoradiography. The aHSPGs have then been localized on ovary cryosections taken along gonadotropin-stimulated cycles in immature rats to show the modulation of aHSPG expression in follicles from the ovulatory cohort. In parallel, heparan sulfate core proteins expression was assessed at the mRNA level to clarify the identity of the proteoglycans involved in this system.
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Results |
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We next examined the partition between cell-bound and soluble aHSPGs using normalized values, and Table I shows that 83% aHSPGs were present on the cell surfaces and 17% in the medium under basal conditions. In FSH-primed granulosa cells, aHSPGs were evenly distributed between the cell layer (59%) and the medium (41%), and after LH stimulation a 36% decrease was observed in the cell-bound aHSPGs, which were released into the medium. Statistical analysis showed highly significant differences between stimulated and nonstimulated conditions for soluble and cell-bound aHSPGs (Student paired t-test, control versus FSH, p < 0.05; control versus LH, p < 0.01). These results demonstrate that granulosa cells respond to FSH and furthermore to LH by increasing their aHSPG output, thereby altering their distribution of aHSPGs between cell surface and culture medium in favor of the liberation of cell-bound aHSPGs.
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Expression of HSPG core proteins mRNA in the rat ovary and in isolated granulosa cells
To identify the nature of the HSPG core proteins expressed in the ovary, in particular by granulosa cells, we performed Northern blot analysis on RNA extracted from rat ovaries and from isolated granulosa cells at different stages of the cycle.
We first assessed the expression of HSPG core proteins in ovary extracts and compared their level of mRNA to that in control organs, known for abundant or low expression of these core proteins (Figure 5). The mRNA of five distinct HSPG core proteins were found in the ovary; the basement membrane HSPG perlecan; the membrane-spanning HSPG syndecan-1, syndecan-2, and syndecan-4; and the GPI-anchored HSPG glypican-1. Perlecan, syndecan-1, and glypican-1 mRNA were readily detected; their level, normalized by comparison with glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA, is comparable to that of highly expressing organs. Perlecan mRNA is particularly abundant in the ovary, with a signal stronger than for the kidney, which is a rich source of perlecan. Syndecan-1 mRNA expression in the ovary is comparable with that found in the liver and the brain, with a conserved ratio (3:1) between the two mRNA splicing variants. The signal obtained for glypican-1 mRNA in the ovary is high, representing about 40% of that seen in the brain, which is the major source of glypican-1. Low level of syndecan-2 mRNA was detected in the ovary, similar to the brain and kidney, with comparable expression of the three characteristic splicing variants. Syndecan-4 expression was found at low levels in all organs, consistent with its ubiquitous distribution, with an mRNA content in the ovary comparable to the levels seen in brain and kidney and lower than in the liver.
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Discussion |
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The growth and differentiation of ovarian follicles is largely dependent on sequential stimulation by FSH and LH; FSH induces follicular development and maturation, and LH triggers ovulation and luteinization of the follicle to develop a corpus luteum. We have previously studied the modulation of aHSPG expression in rat granulosa cells stimulated by FSH (Hosseini et al., 1996). The present study extends these observations to subsequent LH stimulation, thereby mimicking the differentiation stage achieved by granulosa cells in vivo at ovulation. We have found that FSH and LH reinforce each other to up-regulate granulosa cell biosynthesis of aHSPGs and to increase the fraction of aHSPGs released in soluble form in the culture medium. The general biosynthesis of HSPG in granulosa cells is known to be increased after gonadotropin stimulations (Adashi et al., 1986
; Ax and Ryan, 1979
; Salustri et al., 1989
; Yanagishita et al., 1981
). However, aHS chains represent about 7% of the total HS synthesized by granulosa cells (Hosseini et al., 1996
), and their production depends on the coordinated action of several biosynthetic enzymes, directing the construction of the pentasaccharide sequence specifically binding AT. In particular, D-glucosaminyl-3-O-sulfotransferase-1 is a key enzyme in aHS synthesis, and its expression level has been directly correlated to aHS synthesis (Shworak et al., 1997
; Zhang et al., 1998
). A similar mechanism could up-regulate aHSPG production by ovarian granulosa cells during gonadotropin-induced differentiation, in parallel to the modulation of the general biosynthesis of HSPGs.
We next set out to evaluate the pattern of expression of aHSPGs during the cyclic development of ovarian follicles in vivo. We have established a method of detection based on the specific binding of aHSPGs by 125I-AT, revealed by microscopic autoradiography. The aHSPGs could be visualized on isolated cultured granulosa cells and on ovary cryosections, and the specificity of aHSPG labeling was demonstrated using glycosidase digestions and glycosaminoglycan competitors. The aHSPG labeling was seen on granulosa cells of antral follicles and on blood vessels endothelial cells, but not on theca cells. The inner follicular compartment remains avascular until ovulation (Amsterdam and Rotmensch, 1987), hence the intense labeling of granulosa cells from large antral follicles is due to aHSPGs endogenous to the follicle, independently from endothelial cell labeling.
We defined the localization of aHSPGs throughout the ovarian cycle induced by gonadotropin treatment of immature animals. Interestingly, primordial follicles, (diameter 1928 µm) containing 8 to 12 flattened granulosa cells in cross sections with no distinguishable theca cells (Hirshfield, 1991), are strongly labeled for aHSPGs at their periphery, and, by virtue of their ability to activate serine protease inhibitors, they might serve to protect primordial follicles from the active proteolysis occurring in neighboring developing follicles. The aHSPGs quickly disappear from follicles at the onset of development in primary and secondary preantral follicles. This down-regulation of aHSPGs is reminiscent of the observed decrease in syndecan-1 expression in migrating keratinocytes during wound healing or in cells with high metastatic potential (Matsumoto et al., 1997
; Subramanian et al., 1997
). The stimulation of animals with DES mimicks the initial stages of follicular development, brought about by the elevated estrogen levels occurring in the preceding cycle. Granulosa cells from DES-treated animals are weakly labeled for aHSPGs, indicating that estrogens alone do not induce aHSPGs. In contrast, after FSH induction, granulosa cells from large antral follicles are heavily labeled for aHSPGs, and this labeling is consistently found throughout the periovulatory period. These observations are in agreement with the increased production of aHSPGs induced by gonadotropins in cultured granulosa cells. Stimulated granulosa cells released increased amounts of aHSPGs in soluble form. In localization experiments, soluble aHSPGs cannot be detected, because the labeling procedure is performed on unfixed material, resulting in the loss of soluble species in washes. In addition, cells in culture are widely exposed to the culture medium and constitutively release increased amounts of soluble material, including proteoglycans, while in tissues the tridimensional architecture provides extensive cellcell and cellmatrix interactions (Kim et al., 1994
). Therefore, the increased labeling of aHSPGs on granulosa cells of preovulatory follicles does not preclude the parallel release of aHSPGs into follicular fluid. Indeed, we have detected aHSPGs in rat follicular fluid, collected during the preparation of granulosa cells from animals stimulated by gonadotropins (data not shown), and large amounts of HSPGs have been detected in human follicular fluid (Eriksen et al., 1999
). These data suggest that aHSPGs accumulate in follicular fluid and are released at ovulation. After ovulation, aHSPG labeling is decreased but still present on luteinized granulosa cells. The follicular architecture is rapidly evolving during corpus luteum formation, the antral cavity fills with a fibrin clot, and granulosa and theca luteal cells intermix to invade this space, closely followed by rapidly expanding capillaries (Fortune, 1994
; Kamat et al., 1995
; Phillips et al., 1990
; Reynolds et al., 1992
). aHSPG staining of corpus luteum is a superposition of a diffused weak yet positive staining of granulosa luteal cells and of the focused labeling of capillaries.
In summary, aHSPGs are very strongly expressed on granulosa cells of ovulatory follicles until ovulation. Hence, we postulate that the inner follicle displays a strongly anticoagulant surface protecting it from fibrin deposition, which ensures the maintenance of fluidity of follicular fluid until expulsion of the oocyte at ovulation. The widespread localization of aHSPGs on the entire granulosa cells volume, and likely also in follicular fluid, is probably necessary to control the large influx of procoagulant plasma proteins that leak from blood vessels during the inflammatory vascular permeabilization occurring at ovulation (Dvorak et al., 1999). It is noteworthy that fibrin depositions occur in outer theca layers, which are devoid of aHSPGs, underlining their functional importance on granulosa cells in the inner follicle. The rapid decrease in aHSPGs observed on the surface of luteinized granulosa cells after ovulation corresponds to a change to a more procoagulant surface, which is permissive for a fibrin clot to form, filling the antral cavity with a provisional matrix for invading luteal cells. Alternatively, aHSPGs could interact with other heparin-activated serine protease inhibitors present in the follicle, such as protease nexin-1 and plasminogen activator inhibitor-1, and be involved in the control of the proteolytic breakdown of the follicular wall at ovulation.
To further analyze the nature of aHSPGs in the follicle, we investigated the identity of HSPG core proteins expressed in the ovary by granulosa cells, to which aHS can be attached. We have identified five HSPG core proteins mRNA expressed by ovarian granulosa cells. The basement membrane HSPG perlecan is present in granulosa cells together with at least four membrane-associated HSPG core proteins, the membrane-spanning syndecan-1, -2, and -4 and the GPI-anchored HSPG glypican-1. Early studies by Yanagishita and Hascall described the synthesis by rat granulosa cells of two HSPG membrane-bound species, one membrane-spanning and one GPI-anchored HSPG core proteins (Yanagishita and Hascall, 1992). Based on the apparent Mr reported for these HSPGs, we propose that they can be accounted for by syndecan-1 and glypican-1. Recently, Ishiguro et al. (1999)
reported the synthesis of the four members of the syndecan family in mouse postovulatory follicles. They report levels of syndecan-3 expression lower than that of syndecan-4 in follicles before ovulation and suggest that syndecan-3 might be involved in angiogenesis in corpora lutea. By analogy, we assume that low levels of syndecan-3 could also be synthesized by rat granulosa cells. In addition, Veugelers et al. (1999)
recently reported that glypican-6 is strongly expressed in the human ovary; however, in situ hybridization on mouse ovary showed mainly glypican-6 expression on mesenchymal cells. Granulosa cells are epithelial cells (Rodgers et al., 1999
), a cell lineage that often contains glypican-1 expression, unlike glypican-6, which is mainly expressed in mesenchymal cells. Moreover, Eriksen et al. (1999)
purified HSPGs in human follicular fluid that are immunologically related but not identical to perlecan, suggesting that additional HPSG might be also present. Hence, rat ovarian granulosa cells express multiple HSPG core proteins. The core proteins target HS chains to their strategic positions on the cell surface or in the extracellular matrix, and they determine their accumulation and turnover at these sites. It remains to be determined if aHS chains are attached to all core proteins synthesized by granulosa cells. Whether cells can specifically direct the attachment of HS chains with particular biological activities on given HSPG core proteins is not firmly established, but mounting evidence suggests some degree of specificity in HS matching to core proteins. aHSPGs have been found on all HSPG core proteins synthesized by endothelial cells, with some preference for glypican (Mertens et al., 1992
). Moreover, it has been recently shown that cells can direct the attachment of HS chains with antiproliferative and adhesion properties to syndecan-1 and glypican-1, respectively (Liu et al., 1998
).
In view of the important variation observed in aHSPG production during the ovarian cycle, we analyzed the expression of perlecan; syndecan-1, -2, and -4; and glypican-1 in pre-ovulatory and postovulatory stages. In total ovary extracts from naturally cycling adult rats, we did not see any striking differences in the signal obtained for HSPG core proteins in pro-estrus, estrus, or metestrus stages, nor did we detect consistent differences in the expression of core proteins mRNAs in granulosa cells isolated from gonadotropin-stimulated ovaries before or after ovulation. Similar observations were reported for syndecan-4 expression in mouse granulosa cells, except that syndecan-4 was up-regulated during apoptosis of atretic follicles (Ishiguro et al., 1999). Granulosa cells from gonadotropin-stimulated ovaries contain little apoptotic cells until ovulation, due to the rescue action of FSH (Chun et al., 1996
). We observed low syndecan-4 mRNA expression before ovulation, close to our limit of detection, consistent with the low expression reported in healthy mice follicles. After ovulation, the syndecan-4 signal was also slightly higher in granulosa cells, probably due to the presence of small atretic follicles that failed to develop fully. This small increase is explained by the minor contribution of apoptotic granulosa cells in the cell population, which is dominated by healthy luteinizing granulosa cells from large postovulatory follicles (data not shown).
Thus, the remarkably stable pattern of mRNA expression of HSPG core proteins in granulosa cells before and after ovulation does not reflect the extensive variations in aHSPGs observed on granulosa cells both in vitro and in vivo. These data indicate that the expression of aHSPGs on rat granulosa cells is regulated at levels distinct from the transcription of HSPG core proteins. Additional modulation could be exerted at the level of the core proteins translation or by alterations of their catabolic pathways (Yanagishita, 1992), including the rapid removal of HSPGs from the cell surface by shedding, a regulatory mechanism that releases intact extracellular domains of the syndecan family members (Subramanian et al., 1997
). Such regulation could explain the reported increase in total HSPG output by stimulated granulosa cells (Salustri et al., 1989
; Yanagishita et al., 1981
), while the mRNA levels of core proteins remain stable. Furthermore, aHSPGs can also be modulated by posttranslational modifications, in agreement with observations by Shworak et al. (1994)
that syndecan-4 core protein was not the limiting factor in aHS chains production. Moreover, the expression of D-glucosaminyl-3-O-sulfotransferase-1 and the assembly of adequate aHS chain precursors have been shown to be limiting factors in the biosynthesis of aHSPGs (Shworak et al., 1996
; Zhang et al., 1999
), and these factors are likely to be involved in the hormonal modulation of aHSPG expression in the ovary. Our observation that D-glucosaminyl-3-O-sulfotransferase-1 is expressed in granulosa cells supports this view.
The ovarian follicle constitutes the only known example of aHSPG expression outside of the vascular bed, and the complex biosynthetic pathway necessary to generate AT-binding sequences suggests that aHSPGs play a key role in the regulation of proteolytic activities in this extravascular compartment. The present demonstration of the highly regulated expression of aHSPGs according to the follicular cycle strengthens the notion that aHSPG function is not restricted to their antithrombotic properties inside the vascular bed. Further studies are needed to identify aHSPG functional partners and to reveal their physiological functions in the reproductive tract.
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Materials and methods |
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Heparan sulfate core protein probes
The plasmid pBR322-BPG5 containing rat perlecan cDNA was generously provided by John Hassell (Shriners Hospital, Tampa, FL) (Noonan et al., 1988). We subcloned the probe prPer01, a 583-bp EcoRISacII fragment of BPG5 rat perlecan cDNA, into pBluescript-KS. Probes for syndecan-1 (pNWS127) and syndecan-4 (pNWS126) were gifts from Robert D. Rosenberg (MIT, Cambridge, MA). Probe pNWS127 is a 585-bp blunt-BsaHI fragment of rat syndecan cDNA inserted into pBluescript-KS. Probe pNWS126 is a 228-bp AccIEcoRV fragment of rat syndecan-4 cDNA inserted into pBluescript-KS (Kojima et al., 1992
). The probe for syndecan-2 (C17) was kindly provided by John T. Gallagher (University of Manchester, GB) (Pierce et al., 1992
). Probe C17 is a 2153-bp full-length cDNA of rat syndecan-2 digested by EcoR1 and inserted into pBluescript-SK. Probes for glypican-1 (4X1 and 4P2) were kindly provided by Arthur Lander (University of Irvine, CA) (Litwack et al., 1994
). Probe 4X1 is a 162-bp EcoRIXhoI fragment of rat glypican (coding region) cDNA inserted into pBluescript-SK (Stratagene). Probe 4P2 is a 332-bp PstI fragment of rat glypican (3' untranslated region) cDNA inserted into pBluescript-SK. The plasmid pmGAPDH.FL, containing the complete cDNA of mouse GAPDH amplified by PCR and inserted into pBluescript-KS, was a gift from Pierre-Alain Menoud (Institute of Histology, Fribourg, Switzerland).
RNA extraction and Northern analyses
Total RNA was isolated from homogenized rat tissues and primary granulosa cells by the guanidium isothiocyanate method as described by Chomczynski and Sacchi (Chomczynski and Sacchi, 1987). The extract was buffered by 0.1 volume sodium acetate 2 M and extracted once with phenol/chloroform/isoamyl alcohol (25:24:1). RNA were precipitated with an equal volume of isopropanol for 1 h at 20°C and recovered by centrifugation. Pellets were resuspended in sodium isothiocyanate 4 M, sodium citrate 25 mM, sodium-sarcosyl 0.5%, and ß-mercaptoethanol and admixed with 1 volume of isopropanol. After centrifugation, pellets were washed with 75% ethanol, resuspended in RNase-free water, and their concentration determined by optical density. RNA samples were denatured in 10 mM sodium phosphate buffer, pH 6.8, containing 1 M glyoxal and 50% dimethyl sulfoxide and resolved on 1.2% agarose gels. RNA samples were transferred to Hybond-N membrane (Amersham) by capillary blotting. The membranes were vacuum backed for 2 h at 80°C. RNA size markers (Promega) were used as size and transfer efficiency markers. For quantitative comparisons of mRNA levels, hybridization signals were compared between the tested mRNA and the housekeeping gene GAPDH mRNA (Belin, 1997
).
Northern blot analysis
Perlecan.
Prehybridization, hybridization, and washes were carried out according to the method described by Nikkari et al. (1994). cDNA probe prPer01 was labeled with 32P-dCTP by using the Prime-a-Gene labeling system (Promega) and hybridized at 2 x 106 c.p.m./ml.
Syndecan-1.
Prehybridization and hybridization were carried out in 0.25 M sodium phosphate buffer, pH 7.2, containing 7% SDS, 1 mM EDTA, and 1% BSA. pNWS127 cDNA probe was labeled with 32P-dCTP with the Prime-a-Gene labeling system and hybridized at 1 x 106 c.p.m./ml. Hybridization was carried out at 48°C for 1520 h. The blots were then washed twice at 48°C in 2% SDS, 0.1 M sodium phosphate buffer, 1 mM EDTA for 15 min.
Syndecan-2.
Prehybridization, hybridization, and washes were carried out according to the method described by Pierce et al. (1992). The C17 probe was labeled with 32P-dCTP using the Prime-a-Gene labeling system and hybridized at 2 x 106 c.p.m./ml.
Syndecan-4
. Prehybridization, hybridization, and washes were carried out according to published procedures (Belin, 1997). pNWS126 plasmid was linearized by XbaI, and pNWS126 cRNA probe was labeled with 32P-UTP using T3 polymerase. The probe was hybridized at 2 x 106 c.p.m./ml.
Glypican-1.
Prehybridization, hybridization and washes were carried out according to the method described by Litwack et al. (1994). 4X1 and 4P2 cDNA probes were labeled with 32P-dCTP and 32P-dATP (Hartmann, Switzerland) using the Prime-a-Gene labeling system and hybridized at 2 x 106 c.p.m./ml. The two probes were hybridized simultaneously.
All radiolabeled probes were separated from unincorporated nucleotide using G-50 Sephadex home-made 1-ml columns. All filters were exposed to Kodak XAR-5 film with two intensifying screens at 80°C for 2448 h. Quantification of hybridization signal intensity was performed using a Phosphorimager (Molecular Dynamics, Sunnyvale, CA).
RT-PCR detection of D-glucosaminyl-3-O-sulfotransferase-1
First-strand cDNA was generated in a 20 µl volume from 1 µg total RNA of different organs primed with gene specific primer (3A rat primer : 5'-TTGGGATCTACTTGAGGGTGCGC-3', 1 pM) using a reverse transcriptase (RT, Life technologies) according to manufacturers protocol. Touchdown PCR reaction conditions (10 µl) were carried out according to the method described by Shworak et al. (1997), using 1S rat primer: 5'-CCTGGCCCGGGACTCAAACAGCAGGG-3' and 2A rat primer: 5'-TCCCAGTCAAAGAAATGGACCTCGTTTTC-3' (rat cDNA sequence accession number AF177430), giving a 219-bp band resolved on 2% agarose gel electrophoresis with SYBR green staining (Molecular Probes). Rat microvascular endothelial cells (RFP), used as positive control, were described previously (de Agostini et al., 1994
).
Animals
Immature 23-day-old female Sprague-Dawley rats, purchased from Iffa-Credo (LArbresle, France) were maintained on a 12 h day/night cycle and treated with gonadotropins to induce ovulation by sequential injection of PMSG (20 IU) and hCG (10 IU) 48 h later according to published procedures (Peng et al., 1993). The animals were sacrificed by decapitation, and the ovaries were recovered before treatment (immature control), 48 h after PMSG injection, and 6 h, 12 h, 24 h, or 72 h after hCG injection. Alternatively, gonadotropin-independent granulosa cell proliferation was induced by treating 21-day-old female rats by daily subcutaneous injections of 1 mg DES in sesame oil for 4 days; the animals were sacrificed at day 25, and ovaries were used to isolate granulosa cells for culture or were frozen for microscopic analysis (Hosseini et al., 1996
). Regular cycles in adult female rats were documented by daily vaginal cytology for 10 days. Animals were sacrificed, and ovaries recovered at proestrus (preovulatory phase), estrus (ovulation), and metestrus (postovulatry phase).
Granulosa cell culture
Ovaries from DES-treated animals were dissected, granulosa cells isolated as described (Hosseini et al., 1996), seeded in parallel in 96-well plates and on microscope slides precoated with 1 mg/ml bovine vitronectin at 0.5 x 106 cells/cm2 and cultured for 48 h in McCoys medium supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mM L-glutamine, 0.03 mM isobutylmethylxanthine, 2.6 mM 19-hydroxyandrostendione, and 50 ng/ml ovine FSH in humidified 5% CO2 /95% air at 37°C. For LH stimulation, the FSH-primed cells were subsequently incubated for an additional 48 h with fresh medium containing 0.5 µg/ml ovine LH. The response of cultured granulosa cells to FSH and LH was verified by quantifying estradiol and progesterone in spent medium by RIA, respectively (data not shown). Cell-bound and secreted aHSPGs were measured at the cell surface and in the medium by 125I-AT cell- and ligand-binding assays as described (de Agostini et al., 1994
). The protein content of control wells was measured using the BCA protein assay (Pierce), and results were normalized as c.p.m./mg protein. For RNA extraction, granulosa cells were obtained as outlined above, recovered by centrifugation, and pellets were immediately extracted as described.
Tissues
Ovaries for microscopic localization of aHSPGs were rapidly excised from sacrificed animals, dissected free of ovarian fat pad, and embedded in Tissue Tek OCT compound (Miles Inc., IN, USA), frozen in precooled 2-methyl-butane and stored at 80°C. Five-micrometer cryosections were cut from frozen ovaries using a Microm cryostat, mounted on poly(L-lysine)-coated slides, and stored at 20°C until used. Duplicate serial sections were used for incubations with 125I-AT, histologic staining with hematoxylin/eosin, and immunohistochemistry. For RNA extraction, dissected ovaries were snap-frozen in liquid nitrogen and kept at 80°C until extraction.
aHSPG localization
aHSPGs were localized by 125I-AT-binding on ovary cryosections followed by microscopic autoradiography. Cryosections were air-dried and then preincubated in phosphate-buffered saline (PBS) (NaCl 0.15 M, sodium phosphate buffer 10 mM, pH 7.4) containing 50 µg/ml BSA (Sigma) for 15 min. The sections were incubated with 35 µl of 125I-AT diluted in the same buffer (15,000 c.p.m./µl) for 1 h at 4°C in humidified chambers. Excess unbound 125I-AT was removed by five washes at 4°C, with three changes of PBS containing 100 µg/ml BSA and two changes of PBS. After fixation for 10 min in ethanol at 20°C, the section were rehydrated in 70% ethanol and distilled water. After drying, the slides were immersed in NTB-2 photographic emulsion and developed after 35 or 610 days exposure for dark field and light field illustrations, respectively (Sappino et al., 1989). aHSPG localization on cultured granulosa cells was done using the same technique on live cells. The cells were fixed in ethanol at 20°C after the washes and counterstained in methylene blue for 40 s. The specificity of 125I-AT binding to aHSPGs was shown by competition with soluble sulfated polysaccharides and by preincubation with glycosidases. Control slides were incubated with 125I-AT in the presence of 10 or 100 µg/ml heparin or 100 µg/ml dextran sulfate. Alternatively, control cryosections were preincubated for 1 h at 37°C in PBS containing heparitinase (20 mUI/ml) or chondroitinase ABC (0.1 U/ml) prior to the incubation with 125I-AT.
Endothelial cell staining
Endothelial cells were stained by indirect immunohistochemistry using the specific rat endothelial cell-specific monoclonal antibody (RECA-1) and a peroxidase-conjugated sheep anti-mouse second antibody (Amersham) and revealed with diaminobenzidine as described (Yanagishita et al., 1989). The sections were counterstained with hematoxilin. When the first antibody was replaced by a control non-immune antibody, no staining was observed (data not shown).
Observation and photography
Stained preparations were examined using a Nikon Optiphot-2 microscope equipped with Nikon E-plan 10/0.25 Ph1DL and E-plan 40/0.65 Ph3DL objectives (Nikon, Japan) for ovary sections and cultured cells, respectively. Micrographs shown in dark field exposure of autoradiogram of 125I-AT-labeled ovary sections were taken with a photo camera system HFX-DX attachment using Kodak Ektachrome film. Micrographs shown in bright field exposure of stained ovary sections were taken using a Zeiss Axiophot photomicroscope (Carl Zeiss) equipped with Plan Neofluar 10/0.30 and Ph2 40/0.75 objectives and with a high-sensitivity Coolview color digital camera (Photonic Science, London, UK). All pictures were compiled by using PhotoShop version 5.0 (Adobe System, Mountain View, CA) and printed with a digital Fujifilm Pictography 4000 printer (Fujifilm, Tokyo, Japan) (Christen et al., 1999).
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Abbreviations |
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Footnotes |
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References |
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