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Endocrinology Vol. 148, No. 8 3730-3739
Copyright © 2007 by The Endocrine Society

In Vitro Evidence Suggests Activin-A May Promote Tissue Remodeling Associated with Human Luteolysis

Michelle Myers, Eva Gay, Alan S. McNeilly, Hamish M. Fraser and W. Colin Duncan

Obstetrics and Gynaecology, Department of Reproductive and Developmental Sciences, University of Edinburgh (M.M., E.G., W.C.D.) and Medical Research Council Human Reproductive Sciences Unit (A.S.M., H.M.F.), Centre for Reproductive Biology, Queen’s Institute of Medical Research, Edinburgh EH16 4TJ, United Kingdom

Address all correspondence and requests for reprints to: Michelle Myers, Obstetrics and Gynaecology, Centre for Reproductive Biology, The Queens Medical Research Institute, 47 Little France Crescent, Edinburgh EH16 4TJ, United Kingdom. E-mail: m.myers{at}sms.ed.ac.uk.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Luteolysis in women is associated with an up-regulation of the expression and activity of matrix metalloproteinase-2 (MMP-2), which is inhibited by human chorionic gonadotropin (hCG) during maternal recognition of pregnancy. Because the primary source of MMP-2 is fibroblasts that do not express LH/hCG receptors, we aimed to investigate the regulation of MMP-2. Women with regular cycles having hysterectomy for nonmalignant conditions and women undergoing oocyte retrieval for assisted conception were used in this current study. Novel primary cultures and cocultures of luteinized granulosa cells and fibroblast-like cells in conjunction with human corpora lutea from different stages of the luteal phase were used to investigate the role of activin-A in the corpus luteum. The effect of hCG, activin-A, and follistatin on MMP-2 activity and expression was assessed by gelatin zymography and quantitative RT-PCR in primary cell cultures. Confirmation of signaling pathways involved in the activation of MMP-2 was assessed by immunofluorescence, RT-PCR, and quantitative RT-PCR. In primary cell culture, steroidogenic cells secrete activin-A and its inhibitors, inhibin-A and follistatin. Follistatin expression is up-regulated by hCG (P < 0.05). The fibroblast-like cells producing MMP-2 have the machinery for activin reception, expressing both type I and type II activin receptors and Smad proteins. Activin-A up-regulated both activity and expression of MMP-2 in fibroblast-like cells (P < 0.05). This activity was inhibited in cocultures of luteinized granulosa cells and fibroblast-like cells in the presence of hCG (P < 0.05) or follistatin (P < 0.01). Activin-A is an excellent candidate for an effector molecule in human luteolysis whose paracrine action is inhibited during maternal recognition of pregnancy.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE HUMAN CORPUS luteum is a highly vascular and active endocrine gland that in the midluteal phase measures up to 2 cm in diameter. However, unless human chorionic gonadotropin (hCG) is secreted by a conceptus, this highly transient structure will undergo functional and structural luteolysis (1), becoming a small fibrous remnant within a matter of days. The process of luteolysis is associated with marked tissue remodeling and vascular involution. Cellular processes that are known to occur during the regression of the human corpus luteum include cell death (2), an increase in the expression of connective tissue growth factor (3), an influx of macrophages (4), and an up-regulation of fibroblast matrix metalloproteinase-2 (MMP-2) expression and activity (5).

MMPs are key proteolytic enzymes involved in the degradation of the extracellular matrix, which is constantly remodeled during luteolysis. Much evidence exists to suggest that MMP-2 is an important luteolytic agent during the demise of the corpus luteum. In rats, structural luteolysis was associated with an increase in MMP-2 activity (6), whereas in pigs, the expression of MMP-2 was elevated in the regressing corpus luteum (7). Additionally, primates (8) and humans (5) show maximal MMP-2 expression in the late luteal phase, positively correlating with the functional and structural regression of the corpus luteum. Furthermore, hCG during maternal recognition of pregnancy in women (5) reduces the expression of MMP-2. Because MMP-2 is up-regulated during luteolysis, and conversely inhibited in the presence of hCG during maternal recognition of pregnancy, it is likely that it has an important role in the luteolytic process.

The primary source of MMP-2 in the human corpus luteum is the luteal fibroblasts that do not express LH/hCG receptors (5), suggesting that there is a paracrine regulator of MMP-2 expression (9). We therefore hypothesized that in the corpus luteum, MMP-2 is regulated by hCG through an intermediate molecule. Unlike LH, FSH, progesterone, estradiol, or inhibin-A, the concentrations of activin-A in serum are maximal during the late luteal phase (10), suggesting it may have a positive luteolytic action. Activins and other members of the TGF-ß superfamily are known to control many diverse physiological processes (11); therefore, we hypothesized that activin-A may be a critical intermediate signaling molecule in the human corpus luteum.

As described previously (4, 5, 12), we have developed a system for the collection of carefully dated human corpora lutea and more recently (3) a primary cell culture system using human luteinized granulosa cells and/or novel cultures of fibroblasts-like cells derived from the luteinizing follicle. The aim of the current study was to investigate whether activin-A was a paracrine regulator of luteal tissue remodeling by investigating its effects on MMP-2 expression and activity using our model systems.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Collection of human corpora lutea
Tissue collection was approved by the local medical research ethics committee, and all women gave informed consent. Human corpora lutea (n = 18) collected over the past 12 yr (3, 5) were enucleated at the time of surgery from women with regular menstrual cycles undergoing hysterectomy for benign conditions and dated on the basis of the urinary LH surge as described previously (9, 13). In this study, six corpora lutea were classified as early-luteal (LH+1 to LH+5), six as mid-luteal (LH+6 to LH+10), and six as late-luteal (LH+11 to LH+14). At operation, the corpus luteum was quartered to ensure that each quarter contained all cellular elements. Two quarters were immediately frozen and stored at –70 C until RNA extraction was carried out. The remaining tissue was fixed in 10% neutral buffered formalin for subsequent immunohistochemistry.

Isolation of human luteinized granulosa cells and derivation of fibroblast-like cells
The medical ethics committee separately approved the collection of cells from patients undergoing assisted conception. With patient consent, follicular fluid was collected from women undergoing transvaginal oocyte retrieval for in vitro fertilization after ovarian stimulation using a standard procedure (14). Isolation of luteinized granulosa cells using Percoll gradient centrifugation was carried out as described previously (3). Fibroblast-like cells were obtained from prolonged cultures of follicular aspirates as described previously (3).

Derivation of fibroblast-like cells from human corpora lutea
Corpora lutea collected during surgery (as described above) were minced in cell culture conditions and placed in flasks containing 10% fetal bovine serum as described previously (3). Cultures were left to reach confluence and grown until sufficient numbers of cells (60,000 per well) could be obtained for experimental procedures described below.

Primary cell culture
Fibroblast-like cells (60,000 per well) were added to 24-well plates. After 24 h in serum-free conditions (supplemented DMEM/F12 Ham mixture; Life Technologies, Inc., Gaithersburg, MD), as described previously (3), fresh medium was added containing human recombinant activin-A (R&D Systems, Inc., Abingdon, UK; 10–100 ng/ml and controls) or inhibin-A (NIBSC, Hertfordshire, UK; 25 ng/ml and control). All controls contained the carrier solution equivalent to the highest concentration added. After 24 h, culture medium was stored for subsequent analysis, and cells were collected for mRNA extraction. Three to four replicates were performed in at least three separate experiments.

Pooled luteinized granulosa cells (100,000 per well) were cultured in 24-well plates precoated with Matrigel (BD Biosciences, Bedford, MA) in serum-free medium (supplemented DMEM/F12 Ham mixture), as described previously (3). Medium was changed every 2–3 d over the course of the culture period. After 6–8 d in culture, a maximally stimulating dose of hCG (Serono Laboratories, Welwyn Garden City, UK; 100 ng/ml and control) was added. After 24 h, medium and cells were stored for subsequent analysis and mRNA extraction. Three experimental replicates were carried out in duplicate.

For coculture experiments, approximately 60,000 fibroblast-like cells were added to the wells containing approximately 100,000 luteinized granulosa cells after culture for 5–7 d. After coculture for 24 h in serum-free medium, fresh serum-free medium was added containing either hCG (100 ng/ml and control) or follistatin (R&D Systems; 500 ng/ml and control). Follistatin treatments were within physiological levels (100–600 ng/ml) of that previously reported in human follicular fluid (15). Each control contained the carrier solution equivalent to the highest concentration added. After 24 h, mRNA was extracted from cocultures and controls consisting of fibroblasts only. Each coculture and control treatment was replicated six times in three separate experiments.

Preparation of cDNA from corpora lutea and cultured cells and primer design
mRNA was batch extracted from both frozen human corpora lutea and cultured cells and reverse transcribed into cDNA using random hexamers as described previously (3). Oligonucleotide PCR primers for each gene investigated were designed using Primer3 software (http://primer3.sourceforge.net/) from DNA sequences obtained from GenBank (www.ncbi.nlm.nih.gov). Primers were synthesized by MWG-AG Biotech (Milton Keynes, UK) and the 5' to 3' sequences used in this study are listed in Table 1Go. PCR conditions were as follows: incubation at 95 C for 5 min, 35 cycles of 95 C for 30 sec, appropriate annealing temperature (Table 1Go) for 30 sec, and 72 C for 90 sec, followed by 72 C for 10 min. PCR products were separated by applying 110 V for 1 h to 2% agarose gels with ethidium bromide and visualized and photographed under UV transillumination.


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  		TABLE 1. Sequence of primers and annealing temperatures used for qualitative and quantitative RT-PCR

 
Quantitative analysis of gene expression by RT-PCR
Each assay was optimized using PCR amplification of human placental cDNA. The assays were optimized for MgCl2 concentrations and annealing temperatures. PCR amplifications were performed using Thermostart Taq (AB Gene, Surrey, UK) in a DNA Engine gradient cycler (MJ Research, Inc., Watertown, MA) as previously described (3). Data were normalized according to the expression level of glucose-6-phosphate dehydrogenase (G6PDH), determined in duplicate by reference to a serial dilution calibration curve generated for each sample using the standard LightCycler software.

Gelatin zymography
Cell culture medium was collected from serum-free cultures and subsequently frozen at –20 C. Aliquots of 200 µl were subjected to freeze drying for 2–3 h until they resembled a powder and then reconstituted in 20 µl sterile dH2O. One microliter of the reconstituted sample was added to sample buffer [10% (vol/vol) glycerol, 1% (wt/vol) SDS, and 0.04% (vol/vol) bromophenol blue] and applied to an 11% (wt/vol) polyacrylamide gel containing 1 mg/ml gelatin and 0.1% (wt/vol) SDS. Gels were incubated in 2.5% Triton X-100 for 45 min after electrophoretic separation of proteins and then incubated at 37 C overnight in digestion buffer [50 mmol/liter Tris-HCl (pH 7.6) containing 0.2 mol/liter NaCl, 5 mmol/liter CaCl2, and 0.02% (wt/vol) Brij 35] as described previously (5). Gels were stained in staining solution [30% (vol/vol) methanol, 10% glacial acetic acid, and 0.5% (wt/vol) Coomassie brilliant blue G250] and then destained in the same solution minus the Coomassie staining dye. The bands on the zymography gels reflect the activity of MMP-2 and were analyzed by transmission densitometry (G-700 densitometer; Bio-Rad, Hemel Hempstead, Hertsfordshire, UK) and integrated software (Quantity One; Bio-Rad). All densitometry measurements were made between samples on the same gel or between gels run under identical conditions with a common control sample on each gel to ensure comparability.

Measurement of inhibin-A and activin-A
Inhibin concentrations in culture medium collected from luteinized granulosa cells were measured using a plate modification of a standard in-house inhibin-A RIA. The sensitivity of this assay was 2 pg/ml and was carried out as previously described (16). Activin-A concentrations were measured using a two-site ELISA kit that measured total activin-A (17) (Oxford Bio-Innovation, Oxfordshire, UK), following the manufacturer’s instructions. This assay had a sensitivity of less than 78 pg/ml and inter- and intraplate coefficients of variation of less than 10%.

Immunohistochemistry
Immunolocalization was carried out using antibodies recognizing phosphorylated Smad 2/3 (New England Biolabs, Hertfordshire, UK), activin ßA subunit (concentration 4.3 µg/ml, kindly provided by Prof. N. Groome, Oxford Brookes University, Oxford, UK), activin receptor IIA (ActRIIA), and activin receptor like-kinase 4 (ALK4) (kindly provided by R. Heldin, Ludwig Institute for Cancer Research, Uppsala, Sweden) in 5-µm paraffin tissue sections of human corpora lutea prepared on poly-L-lysine-coated microscope slides. These sections were dewaxed, rehydrated, and washed in Tris-buffered saline (TBS)/0.1% Tween 20 and TBS, respectively. Sections for phosphorylated Smad 2/3 were subjected to microwave antigen retrieval in 0.01 M citric acid (pH 6.0) for 10 min and left to cool to room temperature. All sections were washed and placed in 3% H2O2/methanol for 30 min to block any endogenous peroxidase activity.

Normal goat serum (NGS; Diagnostics Scotland, Edinburgh, UK) diluted 1:4 in TBS containing 5% BSA (NGS/TBS/BSA) was added to phosphorylated Smad 2/3, ActRIIA, and ALK4 sections, whereas ßA sections were blocked in normal rabbit serum (Diagnostics Scotland) 1:5 in TBS for 1 h at room temperature. Primary antibodies were diluted in respective blocking solutions and incubated on sections overnight at 4 C (phosphorylated Smad 2/3, 1:1000; ßA, 1:500; ActRIIA, 1:400; ALK4, 1:400). Sections were rinsed twice for 5 min each time in wash buffers and incubated with secondary antibodies [for phosphorylated Smad 2/3, ActRIIA, and ALK4, biotinylated goat-antirabbit IgG diluted 1:500 in NGS/TBS/BSA; for ßA, rabbit-antimouse diluted 1:25 in TBS (Dako Corp., Cambridge, UK)].

Incubations lasted for 1 h and were followed by two washes for 5 min. Thereafter, phosphorylated Smad 2/3, ActRIIA, and ALK4 sections were incubated in avidin-biotin complex-horseradish peroxidase (Dako) for 1 h, and ßA sections were incubated in mouse peroxidase-antiperoxidase (Dako) diluted 1:100 in PBS. Incubations were at room temperature for 1 h, and all sections were washed in TBS (twice for 5 min each) and bound antibodies visualized by incubation with liquid 3,3'-diaminobenzidine tetrahydrochloride (Dako). Sections were counterstained lightly with hematoxylin to enable cell identification. Negative controls for each antibody examined were performed identically to the above protocol with primary antibody incubations substituted with blocking serum containing nonspecific Igs at the same concentration. Images were captured using an Olympus Corp. Provis microscope (Olympus Corp. Optical Co., London, UK) equipped with a Kodak DCS330 camera (Eastman Kodak Co., Rochester, NY), stored on an HP computer and assembled using Photoshop 7.0.1 (Adobe Systems Inc., Mountain View, CA).

Double-fluorescent immunohistochemistry
Sections were washed, subjected to antigen retrieval, and blocked as described above. Negative controls were performed as above. Washes detailed below were for 5 min each, and incubations were at room temperature unless otherwise specified. Rabbit anti-phosphorylated Smad 2/3 was diluted 1:100 in NGS/TBS/BSA (see above) and incubated on sections overnight at 4 C. Sections were washed, incubated with goat antirabbit IgG 488 (Dako) diluted 1:200 in PBS for 1 h, and then washed in PBS. Sections were reblocked with NGS/TBS/BSA for 1 h and then incubated with mouse anti-CD31 or mouse anti-CD68 (Dako) diluted 1:20 in NGS/TBS/BSA overnight at 4 C. Sections were washed, incubated with biotinylated goat antimouse IgG (Dako) diluted 1 in 500 in NGS/TBS/BSA for 30 min, and then washed in TBS.

The fluorochrome streptavidin 546 Alexafluor (Molecular Probes, Inc., Eugene, OR) diluted 1:200 in PBS was incubated on slides for 1 h. For the labeling of nucleic acids, sections were counterstained with a nuclear-specific blue fluorescent label (To-Pro 3; Molecular Probes) diluted 1:2000 in PBS for 2 min. Sections were washed and mounted in Permafluor (Beckman Coulter, High Wycombe, UK). Fluorescent images were captured using an LSM 510 Axiovert 100M confocal microscope (Carl Zeiss Ltd., Welwyn Garden City, UK). Images were compiled using Photoshop 7.0.1 (Adobe Systems).

Statistical analysis
Statistical analyses were carried out, after confirmation of normal distributions for parametric analysis, using a paired t test when treatment and control samples were analyzed or with ANOVA when more than two treatments were analyzed. Where significant differences were observed (P < 0.05) using ANOVA, pairwise comparisons were carried out using Bonferroni’s multiple comparisons test. All statistical tests are highlighted in the figure legends, and differences are given as either *, P < 0.05; **, P < 0.01; or ***, P < 0.001. Differences were considered significant at P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
MMP-2 and MMP-9 activity in primary cell culture
MMP activity was examined in primary cell cultures of luteinized granulosa cells and fibroblast-like cells using gelatin zymography (Fig. 1AGo). Primary cultures of luteinized granulosa cells secreted MMP-9, whereas little MMP-2 activity was detected. Conversely, and as anticipated, fibroblast-like cells secreted large amounts of MMP-2. Cocultures of luteinized granulosa cells and fibroblast-like cells secreted both zymogens.


Figure 1
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  		FIG. 1. MMP-2 activity and expression in primary cell cultures. A, Representative gelatin zymogram of primary and cocultures of luteinized granulosa cells and fibroblast-like cells. Luteinized granulosa cells (LGC) secreted MMP-9 and very little MMP-2. Fibroblast-like cells (Fib) secreted only MMP-2 whereas cocultures (Co-cult) of the two cell types show activity for both zymogens. B, Activin-A increases MMP-2 in primary cultures of fibroblast-like cells. Relative MMP-2 mRNA expression, using real-time quantitative RT-PCR, is shown in black bars, and MMP-2 activity, using gelatin zymography, is shown in white bars. MMP-2 expression (P < 0.05, by ANOVA) and activity (P < 0.05, by ANOVA) was significantly increased in fibroblast-like cells exposed to 25 ng/ml activin-A in culture (n = 3–4 experiments). C, Pilot studies using fibroblast-like cells derived from disaggregating human corpora lutea (n = 2) demonstrate the same effect for MMP-2 expression to increase in the presence of 25 ng/ml of activin-A. D, Fibroblast-like cells show mRNA expression for housekeeping gene G6PDH (G6) and MMP-2; however, they do not express the LH receptor (LHR), indicative that the activin action via hCG mechanisms is under paracrine control. Furthermore, the expression of activin receptors and signaling pathways can be confirmed in these cells. Bands represent 200 and 500 bp as indicated. Bgly, ß-Glycan.

 
The effect of activin-A on the expression of MMP-2 in fibroblast-like cells
MMP-2 activity in primary cell cultures was increased by activin-A in a dose-dependent manner (10–100 ng/ml; P < 0.05; r2 = 0.55, by linear regression). Because intrafollicular concentrations of activin-A have been reported to be in the range of 0.42–26.3 ng/ml (18), we investigated the response to physiological concentrations of activin-A (10 and 25 ng/ml) in detail (Fig. 1BGo). Both MMP-2 expression (P < 0.05, by ANOVA) and activity (P < 0.05, by ANOVA) were increased when compared with controls when fibroblast-like cells were exposed to 25 ng/ml for 24 h (Fig. 1BGo). Correspondingly, the primary cultures of fibroblasts expressed activin receptors (I and II) and Smad 2, 3, and 4 as well as MMP-2 (Fig. 1DGo). We believe that the effect of activin-A on fibroblast-like cells is specific because cells exposed to inhibin-A show no change in MMP-2 production (P > 0.05, paired t test, data not shown). Furthermore, pilot studies (n = 2 corpora lutea) on fibroblast-like cells obtained from prolonged cultures of disaggregated human corpora lutea exposed to 25 ng/ml activin-A also demonstrated the same effects on MMP-2 expression (Fig. 1CGo). It is therefore likely that activin-A could act as a regulator of luteal fibroblast MMP-2 production at physiological concentrations.

Identification of activin synthesis and action in human corpora lutea
Corpora lutea, at each stage of the luteal phase, have the potential to synthesize activin-A, respond to activin-A, and inhibit its action because they express mRNA for ßA subunit, activin receptors (I and II), Smad 2 and 3, and the common Smad 4 as well as inhibin {alpha}-subunit, ß-glycan, and follistatin. The activin ßA subunit is localized to the LH-responsive steroidogenic cells of corpora lutea at each stage of the luteal phase (Fig. 2AGo). Although activin receptors can be found on luteal steroidogenic cells, it is notable that luteal fibroblasts, which secrete MMP-2, express both activin receptors I and II (Fig. 2Go, B and C) and nuclear phosphorylated Smad 2/3 (Fig. 2DGo) at each stage of the luteal phase.


Figure 2
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  		FIG. 2. Immunolocalization of factors involved in activin/TGF-ß signaling in human corpora lutea. A, Light-field microscopy of a late-luteal human corpus luteum showing positive brown staining of the ßA-subunit in the granulosa-lutein cells (GLC) with little or none observed in the surrounding fibroblast layer. Inset shows no staining in the negative control serial section. B and C, Positive staining for the activin receptors ALK4 (B) and ActRIIA (C) in the surrounding fibroblast layer of a mid-luteal corpus luteum localized to cells that resemble fibroblast cells (arrows). D, Phosphorylated Smad 2/3 is expressed during all stages of the luteal phase. Positive staining for this receptor-activated Smad is evident in the nucleus of granulosa-lutein cells (GLC), thecal-lutein cells (TLC), and surrounding fibroblast layer (F) of the corpus luteum with little staining around blood vessels (Bv). E, Double immunofluorescence of endothelial cells (CD 31) (arrow) in red colocalized with phosphorylated Smad 2/3 (green) in a mid-luteal corpus luteum. Nuclear phosphorylated Smad staining is clear in presumptive fibroblast cells (asterisk) located in the stromal area of the corpus luteum and less marked in the endothelial cells (arrow). Nuclear staining is depicted in blue. F, High-power images illustrate a low level of phosphorylated Smad 2/3 (green) in some endothelial cells (arrow) in red. G, Low-power image of phosphorylated Smad 2/3 (green) and macrophages (CD 68-positive cells) (red) in a late-luteal corpus luteum. Phosphorylated Smad 2/3 is localized to fibroblast cells (asterisk) and less so in the macrophages (arrow) stained in red. H, Higher-power view of an individual macrophage (red) in the same tissue section (asterisk) showing low levels of phospho-Smad (green). Scale bars, 40 µm (A–D) and 20 µm (E–H).

 
Change in Smad signaling across the luteal phase
We investigated phosphorylated Smad 2/3 across the luteal phase by immunohistochemistry (Fig. 2Go, D–H). Nuclear phosphorylated Smad 2/3 could be detected at all stages of the luteal phase in both the steroidogenic (granulosa lutein cells and theca lutein cells; Fig. 2DGo) and stromal cell compartments. Although it was detected in the fibroblast layer, it was clear that not all cells in this layer immunostained equally. We therefore colocalized phosphorylated Smad 2/3 with other cell markers. Very little phosphorylated Smad 2/3 immunostaining was noted in endothelial cells (CD31-positive cells) when compared with neighboring stromal cells (Fig. 2EGo) at any stage of the luteal phase, although low-level staining could be noted in some cells of vessels (Fig. 2FGo). In all sections examined, the fibroblast layer contained macrophages (CD68-positive cells). Dual immunostaining suggested that there was less phospho-Smad 2/3 immunostaining in the macrophages (Fig. 2Go, G and H) than the fibroblasts.

To investigate changes in Smad signaling across the luteal phase, the expression of Smad 2 and Smad 3, in carefully dated corpora lutea, was investigated using quantitative RT-PCR (Fig. 3Go). Smad 3 expression increased to a maximum in the late luteal phase (Fig. 3BGo) and was different between early and mid (P < 0.05, by ANOVA) and early and late (P < 0.01, by ANOVA) (Fig. 3BGo). The expression of Smad 2 mRNA (Fig. 3AGo) was also assessed over the luteal phase, and although it demonstrated a similar trend to that of Smad 3, the differences did not reach significance.


Figure 3
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  		FIG. 3. Expression of the receptor-activated Smads in human corpora lutea over the luteal cycle. A, Real-time quantitative RT-PCR demonstrated that the mRNA expression of Smad 2 increased over the luteal phase (n = 6 for each group); however, a nonsignificant trend (P > 0.05, by ANOVA) was observed. B, The expression of Smad 3 was significantly increased from the early luteal phase in both mid-luteal (P < 0.05, by ANOVA) and late-luteal (P < 0.01, by ANOVA) samples.

 
Effect of hCG on activin-A and its inhibitors in primary cell cultures of steroidogenic cells
To test our hypothesis that during maternal recognition of pregnancy, the exposure of hCG to the corpus luteum results in less activin-A signaling from the steroidogenic cells, we assessed the effect of hCG in primary cultures of luteinized granulosa cells. The secretion of both inhibin-A (range, 10–500 pg/ml; P < 0.05, by t test) and activin-A (range, 1–5 ng/ml; P < 0.05 by t test) was increased by the addition of hCG within 24 h (Fig. 4Go, A and B). Experimental results were standardized to their controls to consider variation within individual experiments. Although exposure to hCG did not change the inhibin:activin ratio, luteinized granulosa cells markedly increased their expression of the activin-binding protein follistatin (P < 0.01, by t test) in response to hCG stimulation (Fig. 4CGo). In the presence of hCG, the up-regulation of follistatin may therefore be able to reduce the local actions of activin-A.


Figure 4
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  		FIG. 4. The effect of hCG on elements of the activin pathway in primary cell cultures (n = 3). A and B, Luteinized granulosa cells show an increase in inhibin-A (A) (P < 0.05, by t test) and activin-A (B) (P < 0.05, by t test) production when treated with hCG (100 ng/ml) for 24 h relative to control samples. C, The expression of the activin binding protein follistatin (P < 0.01, by t test) is up-regulated in the presence of hCG (100 ng/ml); n = 3 separate experiments.

 
Evidence that luteal MMP-2 is under the paracrine control of activin-A
To test our hypothesis that hCG could inhibit MMP-2 expression in a paracrine signaling fashion, we employed a novel primary cell coculture system of luteinized granulosa cells and fibroblast-like cells (3). In these cocultures, the primary source of MMP-2 in the culture medium are the fibroblast-like cells (Fig. 1AGo). The addition of hCG to fibroblast-like cell cultures had no effect on the expression of MMP-2 (P > 0.05, t test; Fig. 5AGo). However, cocultures exposed to hCG demonstrated a significant reduction in the expression of MMP-2 (P < 0.05, by t test; Fig. 5AGo), suggestive that hCG is acting on luteinized granulosa cells to influence fibroblast-like cell MMP-2 expression in a paracrine manner.


Figure 5
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  		FIG. 5. In vitro model of paracrine signaling in coculture experiments of luteinized granulosa cells and luteal fibroblast-like cells. A, MMP-2 expression was significantly decreased (P < 0.05, by t test) in primary cocultures exposed to hCG (100 ng/ml) for 24 h; B, similarly, the expression of MMP-2 was significantly reduced (P < 0.01, by t test) in cocultures treated with follistatin (FS) (500 ng/ml) for 24 h. Fibroblast-like cell cultures in both treatment groups were unaffected by either hCG or follistatin (P > 0.05, t test).

 
To test our hypothesis that hCG is manipulating MMP-2 expression through a reduction in activin-A activity secondary to an up-regulation of follistatin expression, we added physiological concentrations of follistatin to our primary cell cultures. The addition of follistatin to cultures of fibroblast-like cells had no significant effect upon MMP-2 expression (P > 0.05, t test; Fig. 5BGo). However, follistatin had the same effect as hCG in reducing the expression of MMP-2 in cocultures of luteinized granulosa cells and fibroblast-like cells (Fig. 5BGo; P < 0.01, by t test).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
How luteolysis occurs in women and how it is inhibited by hCG during maternal recognition of pregnancy is not yet understood. What is clear, however, is that the inhibition of luteolysis by hCG involves disparate effects on cell types in the corpus luteum that do not express LH/hCG receptors. We have previously shown that hCG regulates luteal fibroblast (3, 5), macrophage (4), and endothelial cell (19) function. This means that paracrine signaling molecules from steroidogenic cells have key roles in the local regulation of luteal cell function. To date, the identity of these molecules has remained largely elusive. Here we have shown for the first time, using a combination of observational studies on human corpora lutea combined with interventional studies using human primary cell culture and coculture models, that activin-A is an excellent candidate molecule for a paracrine regulator of luteal remodeling during luteolysis whose action can be inhibited by hCG during maternal recognition of pregnancy.

It has to be highlighted that these studies used luteinized granulosa cells and fibroblast-like cells from the luteinizing follicle. We are using this coculture model to inform us about the paracrine interactions that occur during luteolysis. It is possible that dispersed cells from human corpora lutea may respond differently at different stages of the luteal phase. Unfortunately, such comprehensive studies using fresh human luteal tissue are almost impossible, and it is likely that analysis of activin effects on remodeling of luteal cells will need further assessment in subhuman species.

Activins belong to the structurally related TGF-ß superfamily that includes inhibins and bone morphogenetic proteins. Members of this family have been shown to have important paracrine regulatory roles in diverse physiological processes (11). Indeed, activin signaling has been shown to be essential in inflammation (20), cell proliferation and apoptosis (21), fetal development (22), and male reproduction (23, 24). In particular, it has been established that activins may have a paracrine role during the normal ovarian cycle (24, 25). Activin can stimulate the proliferation of granulosa cells in small follicles (26) and enhance their expression of FSH receptors and aromatase (24, 27). It appears that one of the roles of activin in the ovary is to stimulate smaller follicles and inhibit luteinization of larger follicles to maintain the follicle in an FSH-responsive state (24, 28).

There is evidence to support a paracrine role for activin in the corpus luteum. In the first instance, the corpus luteum has the mechanism to synthesize and secrete activin. Studies in women (29) and primates (30) have localized inhibin {alpha} and ßA subunit mRNA and protein to the steroidogenic cells of the corpus luteum. Dispersed luteal cells in vitro and the intact corpus luteum in vivo secrete inhibin-A in a regulated manner (31). The same is seen in cultures of human luteinized granulosa cells. In addition, these cells secrete activin-A, and activin-A is found in the follicular fluid at the time of ovulation (18). Indeed, circulating activin concentrations change across the ovarian cycle, with an increase toward the end of the luteal phase (10). As well as expressing the ßA subunit mRNA and protein, it is likely that the granulosa-lutein cells of the corpus luteum secrete activin-A during the luteal phase.

In addition, we have shown that the corpus luteum has the molecular mechanisms to respond to locally produced activins. Activins signal through two types of transmembrane serine/threonine kinase receptor interactions (32) and the cytoplasmic to nuclear translocation of the intracellular Smad proteins (33). Human corpora lutea express both the type I (ALK 2/4) and type II (A) activin receptors as well as components of the Smad (2, 3, and 4) signaling pathway that are induced by activin. Correspondingly, we have localized the receptors and activated nuclear phosphorylated Smad 2/3 to both steroidogenic and stromal cells of the human corpus luteum. Although the nature of the ligand signaling through the Smad 2/3 pathway is not entirely clear, because TGF-ß signals through similar Smads (32, 34), it is likely that the activin signaling cascade is active and that activins do act locally on different cells types in the corpus luteum.

Activin action, however, is highly controlled in physiological systems. It is tightly regulated by various inhibitors at the ligand, receptor, and postreceptor levels (35). Follistatin, a local regulator of activin, controls activin signaling by forming biologically inactive complexes with the ß-subunits of the activin glycoprotein. Suppression of activin-regulated processes is also achieved by the activin antagonist inhibin and its coreceptor ß-glycan. Inhibin opposes activin action by competitively binding to the same site of the type II activin receptor. Additionally, inhibins have also been thought to have interactions with specific and high-affinity receptors that may then activate signal transduction pathways, such as the inhibitory Smads, that can oppose activin action (36). ß-Glycan on the other hand, has a high affinity for inhibin and forms complexes with the type II activin receptor to block the recruitment of the type I receptor (37) that is required to activate the signaling cascade. As well as making activin, the corpus luteum has at least some of the molecular mechanisms required to inhibit activin.

It is not clear whether activin signaling changes across the luteal phase. It is likely there is more activin available to act in the late luteal phase because the increase in circulating activin-A at the time of luteolysis (10) is not seen with regard to its inhibitors, inhibin A (10, 31) and follistatin (38). We were able to immunolocalize nuclear phosphorylated Smad 2/3, which would be detected in the presence of activin signaling, throughout the luteal phase. Although most increases in Smad signaling are through phosphorylation of the proteins, there are also increases in the expression of Smad mRNA in response to ligand activation. We therefore investigated the expression of Smad 2 and 3 mRNA across the luteal phase. Expression of both Smads tended to be maximal in the late-luteal phase. These results are compatible with activin action increasing in the lead up to luteolysis.

The expression of both activin and its local inhibitors is regulated in the corpus luteum. The addition of hCG up-regulates the inhibin {alpha}-subunit and the secretion of inhibin-A from steroidogenic cells in vitro and in vivo (31, 39). We hypothesized that the inhibin:activin ratio would increase in the presence of hCG and tested this using luteinized granulosa cells in culture. We showed a similar stimulation of both activin-A and inhibin-A and no change to the ratio. However, other studies have shown that short-term gonadotropin stimulation initially increased activin-A secretion, but unlike inhibin-A, this diminishes later in a time- and dose-dependent manner (40), and it is possible that hCG increases the inhibin:activin ratio in the longer term. In addition, both bound inactive activin and free active activin were detected by the assay, and it is possible that the ratio of inhibin to active activin changes. This is particularly important because it is clear that follistatin is hormonally regulated. hCG up-regulates follistatin from luteinized granulosa cells in culture, and follistatin concentrations in follicular fluid are hormonally regulated (38). For instance, follistatin is up-regulated in the serum of pregnant women (15). Collectively, these data are suggestive that luteal activin action, although not necessarily secretion, is inhibited by hCG during luteal rescue.

There appears to be a role for increased activin activity during luteolysis. Previously, activin has been shown to inhibit progesterone production by luteal cells in vitro (41), and therefore it may have a role in the changes associated with functional luteolysis. However, here we suggest that activin has a role in the remodeling associated with structural luteolysis. One of the key features associated with luteolysis is the up-regulation of MMP-2 in stromal cells (5, 6, 42). We have shown for the first time that fibroblast-like cell MMP-2 is up-regulated by concentrations of activin (but not inhibin) found inside the human ovary. For clarity, we use the term fibroblast-like cells and stroma to represent the stromal cells that are directly surrounding the steroidogenic cells and invaginating in between them. Furthermore, stromal MMP-2 expression can be inhibited by hCG in a paracrine manner both in vivo (5) and in coculture models in vitro. The effect of hCG on MMP-2 in these cocultures can be replicated by the addition of follistatin at concentrations found within the human ovary. Because hCG up-regulates follistatin, these results suggest that activin-A may be involved locally in the physiological regulation of MMP-2 activity in the corpus luteum.

MMP-2 is not known for being regulated at the level of its expression. Indeed, an examination of the promoter region of the MMP-2 gene shows fewer regulatory sequences than other MMPs (42), consistent with a gene that is normally constantly expressed. However, there are clearly certain circumstances when its expression is regulated. It is increased in fetal membranes during labor (43) and in the endometrium during menstruation (44), and more importantly, it is increased during natural and induced luteolysis in a wide range of different species (5, 6, 42). The factors involved in this regulation are not certain. There has been a suggestion that steroids are involved in its regulation in the endometrium (45) and cytokines such as TNF-{alpha} may be important in the regulation of endometrial fibroblast (46) and bovine luteal (47) MMP-2 expression. Extravillous trophoblasts overexpressing Smad 4 demonstrate a clear up-regulation of MMP-2 production, mimicking the effect of TGF-ß in this particular system (48). The same theory may apply to activin-A in fibroblast cells of the corpus luteum because we know they express the Smad proteins common to both TGF-ß and activin signaling and that activin-A in fibroblast-like cells increases MMP-2 production. Here, we demonstrate that activin-A can up-regulate luteal MMP-2 expression and activity in vitro at physiological concentrations.

There is some evidence, in other systems, that activin may well regulate MMP-2 expression. Activin-A has been found to regulate MMP-2 in both mouse peritoneal macrophages (49) and human decidua (50, 51). In addition, it has been suggested that activin can up-regulate both MMP-9 and MMP-2 expression in cultured human cytotrophoblast cells (50). However, this was thought to be secondary to activin affecting the differentiation status of these cells. Another molecule from luteal fibroblasts with a role in tissue remodeling during luteolysis is connective tissue growth factor (3). We have shown that activin also up-regulates its expression in luteal fibroblast-like cells (3), although this does not appear to be the only regulating factor. It is not clear whether other molecules as well as activin can regulate luteal MMP-2 expression during luteolysis.

If activin-A is involved in regulating tissue remodeling during luteolysis, it is likely to be self-limiting in nature. This is because the cellular source of activin is the steroidogenic cells that undergo programmed cell death when the corpus luteum is cleared from the ovary (1, 2). It is difficult to determine whether observations made in nonprimate species are relevant to women because there are major species differences in the regulation of the corpus luteum. In the mouse, where ovarian activin ßA expression has been conditionally knocked out (52), the ovary contains multiple corpora lutea when examined. Although is it tempting to speculate that luteolysis may be affected, it has to be remembered that inhibin-A will also be knocked out, and this may well result in higher FSH concentrations than would be expected to cause more ovulations.

Observational studies on human corpora lutea inform us that the steroidogenic cells expressing LH/hCG receptors can secrete activin-A, and the stromal cells that express MMP-2 have the molecular machinery to respond to locally produced activin. Such observations, however, do not tell us the roles of activin, and interventional studies are required. It is now not practical to collect enough well characterized human corpora lutea to disaggregate the cells and manipulate these cells in culture, and other model systems have to be used. The culture of luteinized granulosa cells and the derivation of fibroblast-like cells have the potential to replicate the luteal steroidogenic cell/stromal cell interactions in vitro. Although luteinized granulosa cells are LH/hCG responsive and function in primary culture for 14 d, they may differ from mature granulosa-lutein cells. In addition, although the phenotype of the fibroblast-like cells is identical to luteal stromal cells in all the genes we have analyzed (3) and expression patterns of MMP-2 from prolonged cultures of fibroblasts obtained from corpora lutea, there may be differences. Indeed, we have shown that there are at least two types of fibroblast in the human corpus luteum (53). Furthermore, in culture, the cells do not have the local interaction with immune cells and endothelial cells seen in vivo.

Although we accept there are caveats to our model system and indeed, our in vitro cell culture and coculture models may not entirely mimic what is happening within the corpus luteum, such model systems can give valuable insights into mechanistic functions that otherwise may not be dissected. Therefore, although care needs to be used in the interpretation of their findings and used in conjunction with observational studies, we believe we have the most appropriate model to study and manipulate cell-cell interactions in the human corpus luteum.

In conclusion, we suggest that activin-A may have a physiological role in luteolysis, and part of this role is to stimulate luteal MMP-2 expression. We believe that hCG serves to impede activin action (Fig. 6Go), and this will facilitate luteal maintenance by inhibiting luteolysis and allowing the maternal recognition of pregnancy.


Figure 6
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  		FIG. 6. Schematic of the model we propose for the paracrine regulation of MMP-2 expression by hCG in human luteal cells. A, Steroidogenic cells (SC), but not fibroblast-like cells (FC), express the LH/hCG receptor. The fibroblast cells secrete MMP-2 and express activin receptors. B, During the late-luteal phase, increasing activin-A action from the steroidogenic cells up-regulates the expression of MMP-2 from the fibroblast cells. C, In the presence of hCG during maternal recognition of pregnancy, the increase of inhibin-A will block activin binding to its receptors, whereas follistatin will bind and biologically inactivate secreted activin-A, resulting in decreased activin signaling, thus preventing the increased expression of MMP-2. We suggest that activin-A is a paracrine signaling molecule playing a major role in the regulation of luteolysis in women.

 


   Acknowledgments
 
We gratefully acknowledge the following people: Ian Swanson for performing the inhibin-A assay and Sheila MacPherson for her expertise in double immunofluorescence. We thank Prof. Richard Anderson, Dr. Shiona Coutts, and Helen Wilson for help and reagents; Dr. Joo Thong and Dr. Nirmala Mary for the luteinized granulosa cells; and Dr. Mick Rae, Dr. Vincent Bombail, and Prof. Stephen G. Hillier for helpful discussion.


   Footnotes
 
M.M. is supported by an Overseas Research Scholarship Awards Scheme (ORSAS) award, and the work was supported by the Cunningham Trust.

Disclosure Statement: The authors of this manuscript have nothing to declare.

First Published Online May 3, 2007

Abbreviations: ActRIIA, Activin receptor IIA; ALK4, activin receptor like-kinase 4; G6PDH, glucose-6-phosphate dehydrogenase; hCG, human chorionic gonadotropin; MMP-2, matrix metalloproteinase-2; NGS, normal goat serum; TBS, Tris-buffered saline.

Received February 21, 2007.

Accepted for publication April 23, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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