Molecular Human Reproduction, Vol. 6, No. 4, 324-330,
April 2000
© 2000 European Society of Human Reproduction and Embryology
Ovary and oogenesis |
Luteinized human granulosa cells are associated with endogenous basement membrane-like components in culture
Department of Obstetrics and Gynaecology, University of Southampton, Princess Anne Hospital, Coxford Road, Southampton SO16 5YA, UK
Abstract
Human granulosa cells (GC), prepared from follicular aspirates using a non-enzymic method, were maintained in culture on chamber slides in a defined medium without additional attachment factors or extracellular matrix (ECM). In this system, GC clustered to a limited extent and attached only loosely to the substratum necessitating medium replacement through repeated partial changes to avoid cell loss. Using this new culture system, cell size and progesterone production per cell increased, consistent with continuing luteinization. These processes were associated with maintenance and deposition of endogenous ECM components. Thus, pericellular heparan sulphate proteoglycan (HSPG) was clearly visible by immunocytochemistry around the luteinized GC after culture. Also progressive accumulation of laminin (particularly 
2-, ß1- and 
1-subunits) during culture was shown by Western blotting of GC extracts. Small patches of collagen IV, shown to be already present between freshly prepared GC, were maintained in culture. A clear effect of gonadotrophin on the maintenance of progesterone production in culture was paralleled by an apparent increased pericellular deposition of HSPG. To conclude, luteinization and maintenance of the GC-derived layer of the corpus luteum is likely to involve deposition and conservation of pericellular ECM components.
corpus luteum/extracellular matrix/granulosa cells/HSPG/laminin
Introduction
Following ovulation, granulosa cells (GC) undergo a process of luteinization which involves cell enlargement and increased progesterone production. These luteinized GC eventually form the dominant progesterone-producing layer of the corpus luteum (see review by Behrman et al., 1993). There is immunocytochemical evidence that luteal cells of the mouse (Wordinger et al., 1983
) and, more specifically, luteinized GC of the rat (Matsushima et al., 1996) are associated with extracellular matrix (ECM) in the form of basement membrane-like components including collagen IV and laminin [for molecular composition of basement membrane, see Yurchenko and Schittny (1990)]. Also, gene expression leading to collagen IV and laminin production has been identified in bovine corpus luteum (Zhao and Luck, 1995). Recent reviews (Behrman et al., 1993; McIntush and Smith, 1998) have entertained the idea that this ECM plays a pivotal role in the control of luteal function. Indeed, it is interesting to speculate that ECM may be essential for the survival and maintenance of the epithelial-derived luteinized GC in a way analogous to that suggested for the role of basement membrane in the support of mammary epithelial cells (Pullan et al., 1996). Clearly, changes in the stability of the ECM in the corpus luteum [perhaps through alteration of matrix metalloproteinases (MMP) and their inhibitors] could be fundamental in explaining mechanisms leading both to luteal regression, and to luteal `rescue' initiated by human chorionic gonadotrophin (HCG) in early pregnancy (O'Sullivan et al., 1997; McIntush and Smith, 1998).
There is good, direct evidence that bovine pre-ovulatory GC produce basement membrane-like material in culture (Rodgers et al., 1996). In vivo, this would be deposited between the thecal cell and GC layers of the enlarging follicle. After ovulation, the position is less clear. Although cells of the corpus luteum (and these will include GC-derived, theca cell-derived, endothelial cells and other cells) appear to have the capability of synthesizing basement membrane components (Zhao and Luck, 1995), we do not know which cell type (or types) is involved. It is an intriguing possibility that the pre-ovulatory potential of GC for producing basement membrane elements (Rodgers et al., 1996), is carried over into the early luteal phase in order to establish a new ECM now important for developing corpus luteum function.
There is potential for using human cultured GC derived from follicular aspirates as a model system for looking at deposition of ECM by GC in the corpus luteum of the primate. However, previously described culture systems which required exogenous matrix (e.g. Richardson et al., 1992) or the provision of attachment factors (usually in the form of serum in the medium or as a pre-coating) allowing cells to spread out on the culture surface, would be difficult to use for such studies. We now describe a novel method for culturing human GC which avoids the use of exogenous additions for cell attachment, and obliges the GC to utilize cell–matrix interactions endogenously generated within the culture. Results presented recommend the system for studies on matrix production by GC.
Materials and methods
Patients
Follicular aspirates were obtained at ovum collection for IVF according to a procedure approved by our local ethical committee. The treatment protocol, adapted from a previously described method (Jenkins et al., 1991), involved down-regulation of pituitary function with gonadotrophin releasing hormone analogue (nafarelin: 400 µg intranasally twice daily) started in the luteal phase of the preceding cycle. From day 4 of the IVF cycle, 150–600 IU FSH was administered daily. When the leading two follicles had a diameter of >18 mm and serum oestradiol concentration was >300 pmol/l for each follicle >14 mm in diameter, HCG (10 000 IU) was given and oocytes were collected 34 h later under transvaginal ultrasound guidance. Follicles aspirated were >15 mm in diameter.
Granulosa cells
Follicular aspirates and washes were combined for each patient, and granulosa cells prepared by a method previously described (Richardson et al., 1992) adapted to avoid the use of enzymic dispersion. The medium used throughout the preparation and culture was a mixture (50:50) of Ham's F12 and Dulbecco's modified Eagle's medium supplemented with glutamine (2 mmol/l), penicillin (100 000 IU/l), streptomycin (100 mg/l), amphotericin (0.25 mg/l), insulin (6.25 mg/l), transferrin (6.25 mg/l), selenious acid (6.25 µg/l), bovine serum albumin (BSA; 1.25 g/l) and linoleic acid (5.35 mg/l). Briefly, cells were collected by centrifugation and then centrifuged over 45% Percoll (Pharmacia, St Albans, UK) to remove red blood cells. Cells were harvested from the interface, washed twice by centrifugation and resuspension, counted by haemocytometer and then diluted to 5x105 cells/ml.
Aliquots of cell suspension (0.45 ml/well) were added to 8-well Labtek-II chamber slides (Nunc brand, obtained through Merck, Poole, UK) which provided a pre-washed glass surface for cell attachment. Additions of medium, or HCG (to give 100 ng/ml; activity: approx. 14 000 IU/mg; Sigma, Poole, UK) were included to give a total incubate volume of 0.5 ml. Complete removal and replacement of medium was found to dislodge cells from the culture surface. As a result, two methods were developed for changing media. Firstly, for time-course experiments (shown in Figures 1, 2 and 4b), medium was changed every 48 h by a multi-step procedure involving four cycles of partial removal and replacement of medium so that, assuming perfect mixing, levels of media constituents were reduced by 96%. Cells remained completely undisturbed by this procedure. Secondly, for other experiments (Figures 3, 4a and 5), a simpler one-cycle procedure (removal and replacement of 50% of medium) was adopted. For time-course experiments, different sets of cultures were established which were intended for completion at the different time-points. At the allotted time, all the medium was removed for storage at –20°C, and the cells remaining either taken for DNA assay or extracted for Western analysis.
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DNA assay
After medium removal, DNA assay buffer (0.6 ml) was added to each well and cells scraped from the culture surface into the buffer using a pipette tip. Aliquots (0.45 ml) of the original, freshly prepared cell suspension were centrifuged and pellets dispersed in assay buffer. The cell extracts were assayed for DNA (Labarca and Paigen, 1980
) using calf thymus DNA as standard. Compared with levels of DNA detected after culture with HCG, levels following culture without gonadotrophin were reduced by ~30–50% by day 8, probably reflecting poorer cell survival and loss from the culture surface. Similar levels of cell loss were detected using an assay based on neutral red uptake by the cultures which has been used previously as a measure of viable cell number (Campbell et al., 1996).
Progesterone assay
Progesterone was assayed in culture medium using an automated, solid-phase, chemiluminescent enzyme immunoassay (Immunolite Analyser System; Diagnostic Products Corporation, Llanberis, UK). Over a working range of 20–120 nmol/l; the intra- and inter-assay variations were <7% and <12% respectively. Samples from one experiment were always run as one batch. Progesterone production by cells (Figure 1
) was corrected for DNA measured for the time and culture condition shown.
Western analysis
Sample buffer [50 µl; 3% (w/v) sodium dodecyl sulphate (SDS), 10% (v/v) glycerol, 1 mg/ml bromophenol blue, 0.0625 mol/l Tris–HCl (adjusted to pH 6.8)] was added to each well at ~60°C. Cells were scraped off the culture surface with a pipette tip and extracts combined before storage at –80°C. Equivalent wells were taken for DNA assay. Freshly prepared cells were centrifuged, medium removed and hot sample buffer (50 µl for each 0.45 ml of cell suspension) added prior to vigorous mixing and storage as above. Before electrophoresis, aliquots of extracts were reduced with 100 mmol/l dithiothreitol for 2x10 min at 95°C with mixing between heating periods. Aliquots of extracts (~30 µl), volumes standardized for cell content on the basis of DNA assay, were subjected to SDS–polyacrylamide gel electrophoresis using 6% resolving gels. Also run were coloured molecular weight markers (Bio-Rad, Hemel Hempstead, UK), and reduced extracts (see above) of both Matrigel (Stratech, Luton, UK) providing subunits of mouse laminin, and human placental laminin (Life Technologies, Paisley, UK). Proteins were blotted onto polyvinylidene difluoride membranes using the Mini Trans-Blot Cell (Bio-Rad). The membranes were blocked in Tris-buffered saline with Tween [TBST; 0.05 mol/l Tris–HCl, 0.15 mol/l NaCl, 0.1% (v/v) Tween 20, pH 7.5] containing 10% dried milk powder. Blots were then exposed for 2 h at room temperature to primary antibodies dissolved in TBST with 10% milk powder. Antibodies used were: monoclonal antibodies against the 2-subunit (Life Technologies), ß1-subunit (Chemicon, Harrow, UK; MAB1928) and 1-subunit (Chemicon; MAB1914) of laminin, and a polyclonal antibody raised in rabbits against human placental laminin (Autogen Bioclear, Calne, UK). After rinsing in TBST, membranes were incubated for 1 h with the peroxide conjugate of the appropriate secondary antibody (Sigma) and then developed using the ECL-plus system (Amersham Pharmacia Biotech, Little Chalfont, UK). Controls carried out under identical conditions, but without primary antibody, were negative.
Immunocytochemistry
Cultures or freshly prepared cells (allowed to settle) were fixed with 4% (w/v) paraformaldehyde for 15 min, rinsed with Dulbecco's phosphate-buffered saline (PBS) and then exposed to 10% (v/v) normal goat serum in PBS for 30 min. Cells were then exposed to primary antibodies in the 10% goat serum overnight at 4°C. Antibodies used were: monoclonal antibodies against human collagen IV [raised against human glomerular type IV collagen, specificity detailed by Gusterson et al. (1984) and supplied by Novocastra, Newcastle-upon-Tyne, UK] and heparan sulphate proteoglycan (HSPG; 10E4 epitope; AMS Biotechnology, Witney, UK), and the polyclonal anti-human laminin (see above). Control chambers without primary antibody were run in parallel. On the following day, cultures were washed with PBS-A [PBS containing 0.2% (w/v) BSA], exposed to an appropriate rhodamine-labelled F(ab')2 fragment secondary antibody raised in goats (Stratech), rinsed in PBS-A and mounted in Citifluor (Citifluor Products, Canterbury, UK). Preparations were examined using a Leica TCS 4D confocal microscope, with a krypton/argon laser as excitation light source arranged for epifluorescence. Images shown in Figure 3 are `virtual confocal slices' where scanning parameters have been standardized, allowing comparison between images for each ECM component examined.
Results
Figure 1 shows an initial elevation of progesterone production by luteinizing GC in culture irrespective of the presence of gonadotrophin. After day 4 of culture, a falling production of progesterone under control conditions contrasts with an effective maintenance of steroid production in the presence of HCG. The gross morphology of the cultures (Figure 2) shows that some clustering of the cells occurred under both control and hormone-stimulated conditions. However, the appearance of the HCG-maintained clusters, which look like `bunches of grapes', differed from the morphology of those under control conditions where outlines of the cells were less clear and clusters appeared smoother and more rounded in outline.
Immunocytochemistry for the main constituents of basement membrane is shown in Figure 3, both in the freshly prepared GC (Figure 3a,c,f) and in the enlarged GC evident after culture (Figure 3b,d,e,g). These results are also summarized in Table I. Small, irregular patches of collagen IV were present apparently often between cells before and after culture (Figure 3a,b). Within the limits of the technique, there was no apparent difference in the level of collagen IV staining between hormone-treated and control cells (only HCG-treated cells are shown). Immunostaining for HSPG was only faintly visible in the freshly prepared GC (Figure 3c). After culture, amounts of HSPG increased (Figure 3d,e) and were particularly elevated around cells which had been exposed to HCG (Figure 3e) where a continuous pericellular coating was apparent. Immunostaining using the polyclonal antibody against laminin showed highly variable levels of laminin around the smaller, freshly prepared, cells with many showing negligible immunoreactivity (Figure 3f). After culture, the surviving, large GC in clusters showed a thin, but more consistent, pericellular layer of laminin (Figure 3g) which, spread over the much larger surface area of the cell, suggested net deposition of laminin in culture. Within the limits of the technique, there was no noticeable difference in the extent of immunostaining for laminin when HCG-treated and non-treated GC were compared (only HCG-treated cells shown).
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In order to obtain another estimate of laminin synthesis in culture, Western analyses of cell extracts was carried out. Western analysis of a placental laminin standard using the polyclonal antibody against laminin (Figure 4
) revealed a major immunoreactive band at ~200 kDa. This is probably reflecting the level of ß- and/or -subunits of laminin and is likely to provide a good `general' impression of the presence of laminin where knowledge of the exact identities of the various subunits is not available as in this case. The level of this `200 kDa laminin' was higher in the cultured cells than in the freshly prepared cells (Figure 4a) and increased gradually over time in culture (Figure 4b). The lower level of laminin in the original cells (Figure 4a) was a reproducible observation apparent in a total of five preparations examined and was evident whether or not the amount loaded onto the gels was adjusted for cell content on the basis of DNA (adjusted loading, Figure 4a; unadjusted loading, data not shown). Over a range of experiments (e.g. those shown in Figure 4a and b) we did not detect an increased level of laminin deposition following HCG stimulation. Western analysis using monoclonal antibodies against specific laminin subunits is shown in Figure 5. The antibody to the 2-subunit is known to be specific for an 80 kDa fragment which is generated by post-translational proteolysis, non-covalently associated with intact laminin and released during the extraction process. Strong bands of immunoreactivity at ~80 kDa in extracts from the cultured GC (and human placental laminin standard) are therefore consistent with the presence of 2-laminin (Figure 5a). Similarly, blots developed with the antibody against ß1-laminin (Figure 5b) showing strong immunoreactivity at about 200 kDa in extracts of cultured cells and Matrigel, are consistent with the presence of ß1-laminin (n.b. the antibody reacts with both the human and mouse ß1-laminin). Also, there was evidence for the presence of the 1-subunit of laminin (Figure 5c). Like the results obtained with the polyclonal antibody (Figure 4), much lower levels of laminin subunits were found in the extracts from freshly prepared cells irrespective of whether a loading adjustment was made for cell number. Also, levels of the three subunits were not noticeably altered after HCG exposure in culture. Results for the three subunits were consistent over a total of five experiments. Discussion
It is a significant feature of the present study that the GC were not provided with exogenous attachment factors or extracellular matrix in culture. Also, the interaction with the glass surfaces of the Labtek-II chamber slides remained relatively loose. Despite this, the cultured GC showed evidence of completion of luteinization in culture both in terms of increased progesterone production (Figure 1) and cell enlargement to cell diameters consistent with mature granulosa-derived luteal cells obtained from dispersed corpus luteum (Webley et al., 1989). As published evidence shows that cell–matrix interactions may be essential for luteinization (Aten et al., 1995), we speculated that our data contributed circumstantial evidence that the cultures may be in contact with some form of endogenous matrix. Our suspicions were reinforced by evidence of some clustering of the GC in culture (Figure 2), as clustering appears to be a phenomenon associated with the availability of matrix (Richardson et al., 2000). Also, the general features of steroid production, with and without gonadotrophin (Figure 1), were not dissimilar to previous work which utilized exogenous ECM (Richardson et al., 1992). Once again, our results were not inconsistent with the generation of endogenous cell–matrix interactions within the GC cultures.
The observation that deposits of collagen IV are present amongst freshly prepared human GC (Figure 3a) is consistent with a report by Yamada et al. (1999) where collagen IV was demonstrated using flow cytometry and immunocytochemistry. Our work now shows at a higher magnification using confocal microscopy how these small patches of collagen IV are positioned around and between the GC. Although the physiological significance of this collagen is not known, its formation may either be an aspect of the process whereby basement membrane components (intended for the basement membrane between GC and theca cells in the follicle) are synthesized by GC (Rodgers et al., 1996; Zhao and Luck, 1996), or the start of a process associated with luteinization. Our finding that the deposits of collagen IV are, at least, maintained in culture is consistent with the results of Yamada et al. (1999) although their study seemed to indicate that an upregulation of collagen IV production occurred once the cells had been put into culture, and that this was augmented in the presence of HCG as measured by release into the medium. Any potential differences between uncultured and cultured cells, and between cells untreated and treated with HCG, were not of sufficient magnitude in our study to show up clearly using immunohistochemistry. Despite these differences between the present study and that of Yamada et al. (1999), the two studies do agree on the important conclusion that if deposition of collagen IV is part of the luteinization process, this process is well underway in women before ovulation (at least in IVF cycles).
The observation that the luteinized GC acquired a pericellular coating of HSPG (Figure 3d,e) is consistent with previous studies showing the synthesis of HSPG by rat granulosa cells (Yanagishita and Hascall, 1983), and the presence of HSPG around rat luteal cells (Asakai et al., 1993). Although there is evidence for HSPG in human follicular fluid (Eriksen et al., 1997) indicating that HSPG may have an important role in primates, there appears to be little related information hitherto on its cellular production and disposition. The much stronger signal for HSPG after culture of GC (compared with the freshly prepared cells) suggests that considerable extra deposition of HSPG may occur as part of the process of luteinization. This newly formed HSPG would have the potential to bind a range of growth factors, including basic fibroblast growth factor (Asakai et al., 1993), which might be important in maintaining the differentiated function of the luteal cell (Aharoni et al., 1997).
The finding that variable amounts of laminin were present within the same preparation of freshly prepared human granulosa cells (Figure 3f) is entirely consistent with previous work (Fujiwara et al., 1997). After culture, the laminin detected immunohistochemically appeared to be more consistent around the bigger, luteinized cells (Figure 3g) so that, on a per cell basis, it was likely that the amount of laminin had increased. This was confirmed by Western analyses of cell extracts which showed a clear and progressive accumulation of laminin in culture measured using the polyclonal antibody to laminin (Figure 4b). Thus, our observations on cultured cells appear to be mimicking the deposition of laminin that occurs during luteinization in vivo, giving rise to luteal cells surrounded by laminin (Wordinger et al., 1983; Matsushima et al., 1996; Fujiwara et al., 1997). It is possible that the increased synthetic capacity for laminin noted in bovine early corpus luteum (Zhao and Luck, 1995) can now be attributed, at least in part, to the granulosa cell fraction.
Laminin isoforms comprise a family of glycoproteins each with a specific subunit structure of -, ß- and -subunits [see Burgeson et al. (1994) for new nomenclature]. Progress needs to be made in identifying the various laminin isoforms present in the ovary and in establishing how the subunit composition of laminin may change as function changes during the menstrual cycle. In the bovine ovary, there appear to be changes in laminin isoforms during follicular development (vanWezel et al., 1998) and a switch to the production of ß1- and 1-subunits of laminin as the corpus luteum is established (Zhao and Luck, 1995). The present study looks at the laminin which may be deposited around luteinizing GC in women and clearly shows the presence of the 2-, ß1- and 1-subunits (Figure 5). Laminin-2 (merosin) is known to contain these subunits and is present in muscle, peripheral nerve and trophoblast (reviewed by Church et al., 1997). It should be noted, however, that a larger study would need to be carried out in order to exclude the possibility of the presence of other laminins such as laminin-4 (2ß21). Our findings may have implications for the development of cell culture techniques for GC. If we are to provide exogenous ECM in vitro, we may be obliged to supply the cells with the physiological laminin isoform for luteinized GC, rather than the mouse laminin-1 used in previous studies (e.g. Fujiwara et al., 1997).
There were some differences in the results obtained for the three major components of basement membrane studied. It appeared that the 34 h exposure to HCG in vivo was sufficient to trigger an initial rise in progesterone production by GC cultured without gonadotrophin, and that this was sufficient to cause laminin deposition in culture. This is the probable explanation for the lack of effect of HCG in culture on laminin production in our study. These results with laminin contrast with our findings on HSPG production where a reproducible stimulatory effect of HCG in culture was seen (Figure 3e). The characteristics of collagen IV production are different once again in that considerable immunostaining is already apparent in the freshly prepared GC so that any increases in culture are not sufficiently clear to be easily detectable using immunohistochemistry. Furthermore, the deposits of collagen IV show a patchy morphology which is quite different from the more uniform pericellular deposition of laminin and HSPG. Clearly, the major constituents of basement membrane around luteinizing GC show separate and distinct characteristics and may be established according to different timetables.
To conclude, we have established a cell culture model without the use of exogenous ECM or attachment factors which now enables us to examine the deposition of matrix which may occur as GC enlarge to become GC-derived cells of the corpus luteum. How this matrix influences luteinization remains to be established. While some work suggests that ECM may have roles in supporting cell survival and steroidogenesis of GC (Aharoni et al., 1997), other studies (Fujiwara et al., 1997; Yamada et al., 1999) have shown an inhibitory action of exogenous matrix components on luteinization. The present study may help us move towards the use of culture models which are closer to the physiological situation and where cultured GC are in contact with the appropriate ECM. Such an approach may assist in reconciling the different points of view regarding the role of cell–matrix interactions in luteinization.
Acknowledgments
We would like to thank the Wessex Fertility Unit, Princess Anne Hospital for the provision of granulosa cells. Financial support from the Solent Subfertility Trust is gratefully acknowledged. The technical assistance of Emma Townsend and Roger Alston (Biomedical Imaging Unit) is much appreciated.
Notes
1 To whom correspondence should be addressed 
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Submitted on October 5, 1999; accepted on January 20, 2000.
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