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Endocrinology, doi:10.1210/en.2005-1228
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Endocrinology Vol. 147, No. 4 1770-1779
Copyright © 2006 by The Endocrine Society

Endothelin-2 in Ovarian Follicle Rupture

CheMyong Ko, Mary C. Gieske, Linah Al-Alem, YunKyung Hahn, Wen Su, Ming C. Gong, Marc Iglarz and Yongbum Koo

Departments of Clinical Sciences and Biological Sciences (C.K., M.C.Gi., L.A.-A., Y.H.) and Physiology (W.S., M.C.Go.), University of Kentucky, Lexington, Kentucky 40506; Pharmacology and Preclinical Development Actelion Pharmaceuticals Ltd. (M.I.), Allschwil CH-4123, Switzerland; and School of Biotechnology and Biomedical Sciences (Y.K.), Inje University, Kimhae 621-749, South Korea

Address all correspondence and requests for reprints to: Dr. Chemyong Ko, Department of Clinical Sciences, University of Kentucky, Lexington, Kentucky 40536. E-mail: cko2{at}uky.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The ovulatory process is activated by a surge of LH, a pituitary gonadotropin, which initiates a cohort of dramatic changes in biochemical, physical, and gene expression in the ovary, leading to follicle rupture and oocyte release. Here we report the identification of endothelin-2 (EDN2) as a last moment-trigger of follicle rupture. In the ovary, EDN2 is exclusively and transiently expressed in the granulosa cells immediately before ovulation. Administration of EDN2 to the ovarian tissue induced rapid contraction, whereas addition of tezosentan, an endothelin receptor antagonist, diminishes the EDN2 effect. In vivo, treatment of tezosentan before ovulation substantially decreases gonadotropin-induced superovulation. As a target tissue of EDN2 action, we identified a layer of smooth muscle cells in the follicular wall of each follicle. Taken together, our data indicate that EDN2 induces follicular rupture by constricting periovulatory follicles.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
OVULATION, THE expulsion of an egg from a mature follicle, is the culmination of a life-long process of female gamete (egg) production and absolutely critical for perpetuating the survival of a species. Coordinated actions of the multiple hormonal and cellular components of the hypothalamic-anterior pituitary-ovarian axis control this process (1, 2). For the last two decades, enormous efforts have been made to reveal the biophysical and molecular mechanism of ovulation, mostly focusing on the identification of the genes that are induced by the surge of LH, a signal that activates the ovulatory program in the preovulatory ovary. Even though numerous genes have been identified as the mediators of LH action in the ovary, the molecular and physiological mechanisms governing the final moment of ovulation (follicular rupture) is still unknown. Failure in ovulation results in infertility and is responsible for the development of numerous female reproductive diseases such as ovarian cyst development, hormonal imbalance, and polycystic ovarian syndrome (3, 4, 5). In this study, we aimed to identify key genes that are expressed in the ovary and specifically involved in follicle rupture, the final event preceding ovulation. Here, we report that endothelin-2 (EDN2) induced follicular smooth muscle contraction is the driving force of the follicular rupture.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals and reagents
The University of Kentucky Animal Care and Use Committee approved all animal procedures. Immature female C57BL/6 mice or Sprague Dawley rats were acquired from Harlan Sprague Dawley, Inc. (Harlan, IN). The estrogen receptor knockout (ER{alpha}KO) mice were kindly provided by Dr. Pierre Chambon (Institut National de la Santé et de la Recherche Médicale, Paris, France) (6). Animals were maintained in a 14-h light, 10-h dark cycle and given a continuous supply of chow and water. Tezosentan was acquired from ACTELION Pharmaceuticals Ltd. (Allschwil, Switzerland). Antibodies were purchased from Abcam, Inc. (Cambridge, MA). EDN-2 peptide was purchased from American Peptide Co., Inc. (Sunnyvale, CA).

Mouse ovarian gene expression database (mOGED) construction
The tissue collection, RNA extraction, DNA microarray, and database construction were performed as previously described for rOGED construction (7). Briefly, mice were killed at 0 [pre-PMSG (pregnant mare’s serum gonadotropin) injection], 12, and 48 h after PMSG administration or 6, 12, and 24 h after human chorionic gonadotropin (hCG) injection. Upon dissection, ovaries were immediately frozen on dry ice (n = 5 animals/time point) and stored at –80 C for later extraction of total RNA. Total RNA extracted from each time point was pooled together and used for DNA microarray analysis. DNA microarray was performed by the University of Kentucky DNA Microarray Core Facility (Lexington, KY). The DNA microarrays of Affymetrix Mouse Expression Set 430 were used according to manufacturer’s instruction (Affymetrix, Santa Clara, CA).

In situ hybridization
Partial EDN2 cDNA corresponding to EDN-2 mRNA sequence was generated by RT-PCR and used for probe synthesis. Briefly, total RNA (1 µg) isolated from rat preovulatory ovaries obtained at 12 h after hCG injection was reverse-transcribed at 42 C for 1 h using SuperScript II (Invitrogen, Carlsbad, CA) and oligo deoxythymidine primers (Invitrogen). First-strand cDNA samples were amplified using oligonucleotide primer pairs (5'-ggaatgtgtgtacttctgcc-3'and 5'-gcagctcatggtgttatctc-3') designed to amplify the 248- to 628-bp region of the endothelin mRNA (GenBank ID: NM_012549). Amplification consisted of a preincubation at 94 C for 5 min before adding Taq polymerase and then 35 cycles at 94 C for 30 sec, 55 C for 30 sec, and 72 C for 30 sec. A PCR product of the predicted size was cloned into the pCRII-TOPO Vector (Invitrogen). DNA sequences of cloned cDNA determined commercially (MWG Biotech, Inc., High Point, NC).

Ovaries were sectioned at 10-µm on an Ultrapro500 cryostat (Vibratome, St. Louis, MO), mounted onto Superfrost/Plus Microscope slides (VWR, West Chester, PA), and fixed by incubating for 5 min in 5% paraformaldehyde, 5 min in 1x saline sodium citrate (SSC), dipping briefly in water, followed by triethanolamine, then incubating 10 min in triethanolamine/acetic anhydride. After a brief wash in 1x SSC, slides were dehydrated through an ethanol series. Plasmids containing cDNA for the gene of interest were linearized with restriction enzyme, then [{alpha}-35S]uridine triphosphate-labeled RNA probes were synthesized using T7 or Sp6 polymerase. RNA probe (2 x 107 cpm/ml) in hybridization buffer (50% formamide, 0.3 M NaCl, 10 mM Tris, 1 mM EDTA, 1x Denhardt’s reagent, 10% dextran sulfate, 10 mM dithiothreitol, 500 µg/ml polyadenylic acid, and 500 µg/ml yeast tRNA) was applied to sections and incubated at 42 C in a humidity chamber for 15–18 h. Slides were rinsed briefly in 2x SSC, treated with ribonuclease A (RNase A) (20 µg/ml in 2x SSC) at 37 C for 45 min, then taken through a series of washes (2x SSC, 1x SSC, 0.5x SSC, 0.1x SSC) at 55 C for 30 min each, and finally dehydrated through an ethanol series. Slides were exposed to Kodak Biomax XAR film for 4 d and dipped in Kodak NTB-2 emulsion (Eastman Kodak, Rochester, NY) for autoradiography. The slides were exposed for 4–6 wk at 4 C. After developing with Kodak D19, slides were stained with Gill’s no. 1 hematoxylin (Electron Microscopy Sciences, Fort Washington, PA) and 1 mg/ml eosin (Sigma, St. Louis, MO). The signal was visualized on an Olympus CKX41 microscope (Tokyo, Japan).

RT-PCR
A total of 250 ng of random hexamer and 0.5 mM deoxynucleotide triphosphate were added to total RNA (0.5–1 µg) in 12 µl before heating at 65 C for 5 min. Then a cocktail of 1x transcription buffer, 10 mM dithiothreitol, 40 U RNase OUT, and 200 U Moloney leukemia virus reverse transcriptase were added to a total volume of 20 µl. The reaction was incubated for 1.5 h at 42 C, then heated at 72C for 15 min to inactivate RNase H. One microliter of cDNA was added to the 10-µl PCR mixture containing 1x PCR buffer [200 mM Tris-HCl (pH 8.4), 500 mM KCl], 1.5 mM MgCl2, 0.2 mM deoxynucleotide triphosphate, 0.2 µM primer, and Taq DNA polymerase (0.5 U). PCR amplification was done at 20 and 25 cycles on an Eppendorf Mastercycler. The PCR products were separated on a 2.0% agarose gel, stained with SYBR Green I (Sigma) for 20 min, and scanned on a phosphor imager (FujiFilm FLA-5000). Primers were 5'-cca agg agc tcc aga aac ag-3' and 5'-tgg tct ctg tag agt tcc gc-3' for EDN1; 5'-gga atg tgt gta ctt ctg cc-3' and 5'-gca gct cat ggt gtt atc tc-3' for EDN2; 5'-gag gat tgt gtc ccc acc ag-3' and 5'-cgg gtg cag ttt cca act ac-3' for EDN3; 5'-ggc tac aga aga ggc ttg cc-3' and 5'-cat atg cct gcc ctt ccg-3' for L19.

Immunohistochemistry
Sections were cut at 10 µm on a Vibratome cryostat and mounted on Superfrost/Plus Microscope slides (VWR). Immediately after mounting, sections were fixed with 4% paraformaldehyde, and immunostaining was performed using the Vectastain Elite ABC kit (Vector Laboratories, Burlingame, CA). Slides were incubated in 0.3% H2O2 in methanol to destroy endogenous peroxidase activity, then blocked and incubated at 4 C overnight with antibody for smooth muscle actin {alpha} (ab15267) at a dilution of 1:100, endothelin A receptor (ab1919) diluted 1:1000, or endothelin B receptor (ab1923) diluted 1:600. The antigen was detected with the peroxidase substrate diaminobenzidine tetrahydrochloride and nickel chloride. The signal was visualized on an Olympus BX51 microscope.

Isometric tension measurement
Ovaries were dissected at hCG 10 h from the immature rats treated for superovulation. Isometric tension was measured with a force transducer as previously described (8). Briefly, ovaries were placed in a Petri dish containing Kreb’s solution at 37 C. The ovaries were sectioned, and a section of each ovary was mounted onto tissue holder electrodes via silk threads. After equilibrating the sections, they were treated with K+. The ovarian tissues were treated with endothelins followed by tezosentan treatment. The contractility response was monitored and recorded.

In vivo ovulation assay
For the study of endothelin action in vivo, 22-d-old female Sprague Dawley rats were treated for superovulation. Tezosentan was injected 10 h after hCG injection at a dose of 0.5, 5, or 10 mg/kg by ip injection or 0, 01, 1, or 10 ng/kg by intraovarian injection, respectively. The rats were killed and oviduct attached to the ovary was dissected at hCG 20 h for oocyte counting. Oocytes were retrieved from a bulge of an oviduct and microscopically counted. After counting, ovaries were placed on dry ice and stored at –80 C for later histology.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
EDN2 expression in the ovary
We first searched for genes whose expression increased significantly immediately before the rupture of the follicles. A search was conducted using a rat ovarian gene expression database (rOGED: http://app.mc.uky.edu/kolab/rogedendo.aspx), which provides immediate analysis of temporal gene expression profiles for over 28,000 genes in intact ovaries, granulosa cells, and residual ovarian tissues during follicular growth and the periovulatory period (7, 9). This approach followed by RT-PCR confirmation resulted in the identification of EDN2, a potent vasoconstrictor, which was transiently expressed in the granulosa cells of periovulatory follicles (Fig. 1Go, A and B). The in situ hybridization study confirmed that EDN2 mRNA was expressed specifically in the granulosa cells of periovulatory follicles (Fig. 1Go, D and E).


Figure 1
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  		FIG. 1. EDN2 mRNA expression in the rat ovary treated for superovulation. For superovulation, 23-d-old prepubertal rats were injected with PMSG, followed by hCG injection 48 h after PMSG injection. A, EDN2 mRNA expression in the rat ovary, granulosa cells, and residual tissues. This graph was generated by rOGED. The residual ovarian tissues represent mostly theca-interstitial cells (13 ). Error bars are SDs of two independent experiments. B, RT-PCR analysis of EDN2 mRNA expression during periovulatory period. Total RNA pooled from two rats per time point were used for this assay. EDN2 mRNA expression was evident only at hCG 12 h. L19 (ribosomal protein) was used as an internal control. C, External view of rat periovulatory follicles at hCG 12 h. D and E, In situ hybridization was used to localize EDN2 mRNA expression. EDN2 mRNA expression (blue) was evident only in the granulosa cells of hCG 12 h ovary. No EDN2 mRNA expression was detected at nonperiovulatory stages of follicular development. GC, Granulosa cell layer; TIC, theca-interstitial cells; PoF, periovulatory follicle; SaF, small antral follicle.

 
It is necessary for a cell to express endothelin-converting enzyme that catalyzes the conversion of inactive forms of endothelins (163 amino acids) into bioactive endothelins (21 amino acids) to produce bioactive endothelins. A search of rOGED found that endothelin-converting enzyme 1 was expressed in the ovary before ovulation (Fig. 2Go). The search also found the expression of endothelin receptors (ETA and ETB), which are G protein-coupled receptors and convey endothelin function in target cells (10, 11).


Figure 2
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  		FIG. 2. Relative expressions of rat endothelin-converting enzyme 1 (ECE-1) (A), ETA (B), and ETB (C) in intact ovaries, granulosa cells, and residual tissues of the prepubertal rats treated for superovulation. Graphs were generated by the rOGED. Error bars are SDs of two independent experiments.

 
EDN2-induced ovarian tissue contraction
It is well established that the most prominent function of the endothelin receptor system is to constrict smooth muscle tissues (12, 13). Thus, the expression of EDN2 mRNA immediately before ovulation led us to hypothesize that EDN2 may drive the process of follicle rupture by constricting periovulatory follicles. We first evaluated contractile activity of an ovarian tissue in vitro in the presence of EDN2 using an isometric tension measurement approach (8, 14) (Fig. 3Go). EDN2 instantly induced contraction in the ovarian tissue strip obtained from an immature rat primed for superovulation (Fig. 4Go). When added to the contracted tissue, tezosentan, an endothelin receptor antagonist (15), relieved the tissues from the contraction (Fig. 4Go, A and B). This experimental finding demonstrates that the EDN2-induced contraction is modulated by the endothelin receptor-mediated pathway.


Figure 3
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  		FIG. 3. Measurement of follicular contraction. A, Isometric tension measurement system. It is composed of three main components: sensor (left), amplifier (middle), and chart-reader (right). B, High magnification of the sensor component. Two tissue holder electrodes connected to a sensor (bottom) are attached to and controlled by a micromanipulator (top). C, An ovarian tissue strip is held by the two tissue holder electrodes. D, Diagrammatic presentation of the ovarian tissue strip held by the electrodes. Procedure of contraction measurement: ovaries were dissected in a Petri dish containing Kreb’s solution at 37 C. A section of each ovary was mounted onto tissue holder electrodes via silk threads. After equilibrating the section, the tissue was treated with K+. The tissue was then washed and treated with endothelins. The contractile force was then detected by the sensor, amplified by the amplifier, and recorded by a printer.

 

Figure 4
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  		FIG. 4. EDN2-induced follicular contraction. A, Ovarian tissue strips obtained from an hCG 10 h prepubertal rat were treated with 50 nM of EDN2 for 30 min followed by 10 mg/ml tezosentan (TEZO) treatment for an additional 30 min. This is a representative graph for the three independent experiments. B, Summary of three independent experiments are shown. *, P < 0.005 compared with control (K+). C–E, Ovarian smooth muscle system as determined by immunohistochemistry using anti-SMA{alpha}antibody in the hCG 12 h ovary. This is a representative image of four independent experiments. Smooth muscle cells are detected in the theca externa. No positive signal is seen in the granulosa cells (GC) or in the theca interna (TCI). Note that most of the large antral follicles show no SMC{alpha} staining in the apex (red stars). Locations of follicular granulosa layer (GC), basement membrane (BM), theca layer (TC), and smooth muscle cell layer (SMC) are indicated. Blood vessels (BV) are also positively stained (E). F–H, Increased mRNA expression of SMA{alpha} (F), desmin (G), and transgelin (H) before ovulation. Graphs were generated by the rOGED. Note that the mRNA expression is limited to the residual tissues of the ovary, whereas no change in expression level is seen in the granulosa cells.

 
Distribution of smooth muscle tissues in the ovary
Contractile response of ovarian tissue to EDN2 treatment led us to investigate the intraovarian organization of the smooth muscle system in the periovulatory ovary as a target tissue of EDN2 action. Immunohistochemical examination found that individual follicles were surrounded by a smooth muscle layer as shown by the strong immunostaining with anti-SMA{alpha} (smooth muscle actin) in the theca externa of the follicles (Fig. 4CGo). Neighboring smooth muscle layers were directly contacting each other or a layer was shared by an adjacent follicle, forming an interconnected smooth muscle network at the ovarian level (Fig. 4DGo). Interestingly, the apex region of a periovulatory follicle, a putative site of follicle rupture, was not stained by the SMA{alpha} antibody (Fig. 4EGo), indicating absence of smooth muscle tissues in the region. The presence of intraovarian smooth muscle system was further verified by the finding that the periovulatory ovary also expressed other smooth muscle genes such as desmin (a gene encoding a muscle-specific class III intermediate filament) and transgelin (a gene encoding a calponin-related protein that is a marker of contractile activity) (16). The rOGED showed that the expression of desmin and transgelin were specifically detected in the theca cell-enriched residual ovarian tissues and the expression levels increased gradually before ovulation (Fig. 4Go, F–H).

Involvement of ovarian endothelin receptor system in ovulation
We then determined whether the EDN2 receptor system is involved in follicular rupture in vivo. For this purpose, the superovulation regimen was employed (7, 9), in which EDN2 showed maximum expression at 12 h after hCG injection (hCG 12 h) and ovulation occurred between hCG 12 h and 15 h. When two different doses of tezosentan were injected at hCG 10 h by ip injection, a dose-dependent decrease of ovulation was seen (Fig. 5AGo). We then administered tezosentan directly to the medulla area of the ovary by an intraovarian injection to avoid the leakage of tezosentan sometimes observed with the intrabursal method (Fig. 6Go). A dramatic reduction of ovulation was seen in the tezosentan-injected ovary compared with the uninjected (Fig. 5BGo). Furthermore, the ovulation inhibitory effect with 5.0 mg/kg body weight of tezosentan by ip injection was attained by intraovarian injection with a substantially lower dose (10 ng/kg body weight) (Fig. 5Go, A and B). Examination of the ovarian sections revealed that ovaries of the tezosentan-treated rats contained markedly fewer corpora lutea but more unruptured follicles compared with the uninjected ovary (Fig. 5Go, C and D), indicating that the reduced ovulation was due to lack of follicle rupture.


Figure 5
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  		FIG. 5. Inhibition of follicle rupture by the endothelin receptor antagonist, tezosentan. A, Tezosentan treatment by ip injection. Prepubertal rats (26 d old) were treated by the superovulation regimen with PMSG and hCG injections. At hCG 10 h, the rats were additionally injected ip with PBS or tezosentan (TEZO; 0.5, or 5.0 mg/kg body weight). At hCG 20 h, the numbers of released oocytes were counted. The means and error bars (SEM) (PBS, n = 4; TEZO 0.5 mg, n = 5; TEZO 5.0 mg, n = 5) are shown. *, P < 0.005, **P < 0.001 compared with control (PBS). The range of the numbers of released oocytes are indicated in the bars. B, Tezosentan treatment by intraovarian injection. Instead of peritoneal injection, tezosentan was directly injected directly into the medulla of the right ovary (intraovarian) with PBS or tezosentan (TEZO; 10 ng/kg body weight). The uninjected left ovary served as injection control. The means and error bars (SEM) (PBS, n = 10; TEZO 10 ng, n = 7) are shown. A significant reduction of ovulation was found in the tezosentan-injected ovary. *, P < 0.005 compared with control (noninjected). C and D, Histological examination of the PBS-injected (C) and tezosentan-injected ovaries (D). The numbers of ovulated oocytes are indicated in parentheses. Note fewer numbers of corpus lutea (CL) but greater numbers of unruptured follicles (URF) are seen the tezosentan-injected ovary. The numbers of oocytes released from the ovaries (RO) are indicated.

 

Figure 6
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  		FIG. 6. Intrabursal vs. intraovarian injection of tezosentan. Ovaries from a prepubertal rat treated for superovulation were exteriorized surgically at hCG 10 h. Target material was injected either into the space between the ovary and bursa (A, intrabursal injection) or directly into the medullary area of the ovary (B, intraovarian injection). The location of oviduct and fatty tissues are indicated. To show the location of injection in the intrabursal injection, the bursa was intentionally lifted up by the needle (A). After injection, the ovary was placed back into the peritoneal place. Our test injection with a dye solution showed that, during the operation, a significant amount of the injected material leaked out of the ovaries that were injected by intrabursal injection. In contrast no significant leakage was seen in the ovaries injected by the intraovarian injection method. Thus, the intraovarian injection method was employed for the delivery of the tezosentan into the ovary.

 
EDN2 and EDN3 in the ovary
To see whether other isoforms of endothelins were involved in follicle rupture, the contraction inducing activities of EDN1 and EDN3 in ovarian tissue were measured. Although less robust, both EDN1 and EDN3 induced contraction in the ovarian tissue (Fig. 7Go). This finding led us to determine whether these isoforms showed similar expression to EDN2. Investigation of both rat and mouse ovaries revealed that neither isoform displayed comparable expression to EDN2 (Fig. 7Go). It was interesting, however, that endothelin receptors were expressed during the entire period of folliculogenesis both in granulosa cells and residual cells (Fig. 2Go). Consistent with mRNA expression pattern, receptor proteins were detected ubiquitously in the ovary by immunohistochemistry (Fig. 8Go).


Figure 7
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  		FIG. 7. Endothelin isoforms in the ovary. A–C, Relative contraction inducing activities of EDN1, EDN2, and EDN3. Ovarian tissue strips of an hCG 10 h immature rat were treated sequentially with 10, 50, and 100 nM of EDN1, 2, or 3 for 10–20 min, respectively. Note that although the contraction was induced by both EDN1 and 2 over the level of depolarization, EDN3 did not induce contraction higher than the level of depolarization. Shown are the representative patterns of three independent experiments for each isoform. D and E, Relative expression of EDN1 and EDN2 in the rat ovary (D) and mouse ovary (E). EDN3 was not detected in either rat or mouse ovaries (data not shown). The rat profile was generated by rOGED, whereas the mouse graph was drawn by the mouse version of ovarian gene expression database (mOGED), which was generated by the same procedure used for rOGED construction (see Materials and Methods).

 

Figure 8
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  		FIG. 8. Expression of SMA{alpha} (A), ETA (B), and ETB (C) protein in the ovary. Adjacent sections of a hCG 12 rat ovary were analyzed by immunohistochemistry. As was expected from the microarray data, endothelin receptors did not show tissue specificity, but rather showed ubiquitous expression. Not ETA, but ETB showed stronger expression in the smooth muscle layer of follicles (compare the regions pointed by white arrows) and blood vessels (yellow arrow heads) compared with other area of the ovary. Interestingly, ETA showed strong expression in the luminal epithelium (LE) of oviduct. These are representative images of three independent experiments. SM, Oviductal smooth muscle; POF, periovulatory follicle.

 
EDN2 receptor system in the ER{alpha}KO mouse ovary
Current findings indicate that the periovulatory follicle may not rupture if the components of either the endothelin system or ovarian smooth muscle system in the ovary are not functioning properly. As an exemplary case, we examined the hemorrhagic cysts formed in the ER{alpha}KO mouse to see whether EDN2 and/or the smooth muscle system have any causal relationship with cyst development. It is well known that, in the ER{alpha}KO mouse ovary, folliculogenesis proceeds normally up to preovulatory stages but the follicle fails to rupture (17). Thus, we first determined whether the lack of ER{alpha} in the ER{alpha}KO mouse ovary influences EDN2 expression. When treated for superovulation, ER{alpha}KO mice showed a normal level of EDN2 mRNA in the ovary (data not shown), demonstrating that EDN2 expression machinery is intact in the ER{alpha}KO mice. In a comparable manner, immunohistochemistry showed that, although the granulosa layer was severely disorganized, revealing areas of a follicular wall completely devoid of granulosa cells, smooth muscle layers were well maintained (Fig. 9Go). Interestingly, although normal preovulatory follicles lack the muscle layer or have a thinner layer in the apex, the hemorrhagic cyst has a complete circular layer of smooth muscle cells through the wall of the cyst, without showing any thinning or disruption of the smooth muscle layer in any region of the cyst (Fig. 4Go vs. Fig. 9Go).


Figure 9
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  		FIG. 9. Smooth muscle network and EDN2 mRNA expression in the ER{alpha}KO mouse ovary. A, An ovary from a 3-month-old ER{alpha}KO mouse was cut and processed for immunohistochemistry using anti-SMA{alpha} antibody followed by hematoxylin/eosin staining (B and C). The dark black lines in B and C represent SMA{alpha}-positive smooth muscle cell layers (SM). Note the complete round structure of a follicular smooth muscle layer in each of the preantral and antral follicles (PaF, AF) and hemorrhagic cysts (HC). Oviduct (OV) also shows strong immunoreactivity to anti-SMA{alpha} antibody. Although some parts of a follicular wall of a hemorrhagic cyst (HC) are devoid of the granulosa cell layer, the smooth muscle layers are intact (B and C). Note that the smooth muscle layer makes a complete circle in the follicles and hemorrhagic cysts of the ER{alpha}KO mouse ovary, which is in contrast to the incomplete circle in the periovulatory follicles of the wild-type mouse (Fig. 3Go).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Since the introduction of molecular biology into the study of ovulation in the early 1980s, research focus has been centered on the identification of new genes involved in follicle rupture. Accordingly, multiple numbers of genes associated with ovulation have been identified, which has given more weight to the importance of the role of proteolytic activity in follicle rupture. On the contrary, not much attention has been given to the mechanical aspects of follicle rupture. Interestingly, whereas numerous genes have been identified as essential components of follicle rupture, specific roles of the genes in follicle rupture remain largely unknown. For example, whereas progesterone receptors (PRs) and cyclooxygenase-2 (COX-2) are the two most extensively studied LH-induced genes in the ovary (18, 19, 20) and are required for follicle rupture as was shown by the lack of follicle rupture (and the presence of unruptured follicles) in their respective knockout mouse ovaries, the mechanistic roles of the genes in follicle rupture is not known. Furthermore, it is difficult to explain why none of the matrix metalloproteinase (MMP) knockout mice (e.g. MMP-1, -2, -3, -7, -9, -11, -12, and -14 knockouts and tissue inhibitor of metalloproteinase-1, -2, and -3 knockouts) showed major fertility problems (21, 22), even though it was suggested that redundancy in the activity among different MMPs may compensate for the loss of activity in each knockout mouse (21). In addition, whereas many other LH-induced genes have been identified (23), their individual functions in follicle rupture are largely unknown (24). This current difficulty in understanding individual gene functions in follicle rupture demands a new model of follicle rupture. Having a sound model of follicle rupture shall help overcome those presented difficulties in understanding functions of individual genes in the overall context of ovulation and will greatly advance our understanding on the mechanism of follicle rupture at the molecular as well as tissue level.

For decades, it has been speculated that contractile activity may play a key role in follicle rupture. Supporting the speculation, multiple groups reported that follicular pressure increased before rupture (25). However, contrasting findings from different studies and recurring controversy on the reliability of the methods of follicular pressure measurement have diminished the importance of the hypothesis for contractile activity in follicle rupture (26). Nonetheless, a body of evidence from recent findings reiterates the involvement of contractile activity in follicle rupture. For example, it was shown that follicles of hamster and human ovaries contained smooth muscle tissue in the theca externa (27, 28). Other events taking place in the ovary before ovulation include the sharp increase of intraovarian proteinase activity (21, 29), and inflammatory reactions, both of which may undermine the physical integrity of the follicular wall (29). Additionally, excessive extracellular matrix breakdown and cellular death occur before rupture at the apex, or site of rupture (30, 31). We believe that this weakening of the follicle wall may facilitate rupture induced by contraction. The identification of EDN2 as an intrinsic follicular constrictor strongly supports the idea that follicle rupture is driven by follicular constriction.

This study indicates that EDN2 is produced in the granulosa cells but diffuses through the weakened follicular wall, reaching the theca externa where smooth muscle cells are located, and inducing muscle cell contraction (Fig. 10AGo). At the level of a follicle, contraction of individual smooth muscle cells results in the follicular constriction, which increases follicular pressure creating tension in the follicle wall (Fig. 10BGo). Eventually, the follicle will rupture at the apex where the tensile force is weakest due to the lack of smooth muscle cells (Fig. 4Go) and low structural integrity (30, 31) (Fig. 10Go). Because smooth muscle layers are interconnected with one another (Fig. 4Go), the EDN2-induced constriction of individual follicles may generate a synergistic force at the ovarian level, simultaneously rupturing multiple follicles. This may explain why dozens of oocytes are released within a short period of time and travel together in the oviducts in the animals treated by superovulation regime. It is noteworthy that the follicular smooth muscle layer is spatially separated from the site of EDN2 production: EDN2 expression is strictly confined to the granulosa cells of the periovulatory follicle (Fig. 1DGo), whereas smooth muscle is located in the theca externa (Fig. 4Go). It is believed that this physical separation of the site of EDN2 production from its target tissue may be necessary to prevent potential precocious follicular constriction, which could occur if the smooth muscle cell produced EDN2. The basement membrane that is located between granulosa cells and the theca interna (Fig. 10AGo) may serve as an additional physical barrier preventing the free diffusion of EDN2 produced by granulosa cells. Only when the integrity of the basement membrane and the theca interna is disrupted due to the increased proteolytic enzyme activity (21), the EDN2 may easily diffuse through the weakened follicular wall and bind to smooth muscle cells. The narrow temporal window of EDN2 expression at the periovulatory stage, occurring just when the integrity of the follicular wall is disrupted, may also help the follicles to constrict only when they are ready to rupture (Figs. 1Go and 7Go).


Figure 10
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  		FIG. 10. Hypothetical action of EDN2 in follicle rupture (A) LH released from the anterior pituitary travels to ovarian cells inducing a variety of dramatic changes within hours in the ovary: a burst of angiogenesis, infiltration of leukocytes (LC), increased proteinase activity, increase in follicular size, luteinization of granulosa cells (GC), and altered expression of numerous genes. Various proteinases released by leukocytes (red asterisks), theca cells (TI) (blue asterisks), or granulosa cells (green asterisks) weaken the integrity of the follicular wall, whereas the smooth muscle layer (SMC) located in the theca externa maintains the overall structure of the follicle. B, By the time the follicular wall is partially degraded, EDN2 (gray asterisks) produced by the granulosa cells diffuses out of the weakened basement membrane (BM) and theca interna, and easily reaches the smooth muscle cells. The smooth muscle cells then contract (bottom), which eventually constricts the wall of the periovulatory follicle (top). As the contraction force increases (arrows), the apex (the region of the follicular wall facing the surface epithelium of the ovary) is ruptured, through which follicular fluid and the oocytes are released.

 
The smooth muscle distribution in the ER{alpha}KO mouse ovary is of particular interest because no thinning or disruption of the smooth muscle layer in any region of the cyst was seen (Fig. 9Go). Considering the elastic nature of smooth muscle tissues, it is believed that the lack of the apex in the hemorrhagic cyst may make it difficult to rupture. It will be interesting to see whether ER{alpha} plays an important role in the theca externa where the expression of ER{alpha} was reliably detected (32). Furthermore, as PR and COX-2 play critical roles in ovulation as demonstrated by the lack of follicle rupture in the mice deficient in either the PR or COX-2 gene (18, 19, 20, 33), whether these gene are involved in EDN2 receptor system expression or action will be an interesting subject of study.

It is believed that the endothelin receptor system may play a noncontractile role in the ovary because endothelin receptors are expressed not only in the theca externa during the periovulatory period, but also in the granulosa cells and during the nonovulatory phase of folliculogenesis (Figs. 2Go and 8Go). Our preliminary investigation indicates that the endothelin receptor system may be involved in granulosa cell proliferation and survival (data not shown). EDN1 produced by granulosa cells or supplied by circulation (34) may be involved in the noncontractile function of endothelins.

The presence of endothelins in the follicular fluids of women undergoing ovulation induction for in vitro fertilization procedure (35) indicates the use of the endothelin system in human ovulation. Studies on the correlation between ovarian EDN2/smooth muscle system and ovulatory disorders in the human ovary are therefore eagerly awaited.


   Acknowledgments
 
The authors thank Dr. Indrani Bagchi (University of Illinois) for generously sharing the mouse endothelin-2 data she presented at the 37th Annual Meeting of the Society for the Study of Reproduction.


   Footnotes
 
This work was supported by Grants P20 RR15592 and 1RO1HD052694-01 from the National Institutes of Health.

The authors have nothing to declare.

First Published Online January 12, 2006

Abbreviations: COX-2, Cyclooxygenase-2; EDN1–3, endothelin-1, -2, or -3; ER{alpha}KO, estrogen receptor knockout; hCG, human chorionic gonadotropin; MMP, matrix metalloproteinase; mOGED, mouse ovarian gene expression database; PMSG, pregnant mare’s serum gonadotropin; PR, progesterone receptor; rOGED, rat ovarian gene expression database; RNase, ribonuclease; SMA, smooth muscle actin; SSC, saline sodium citrate.

Received September 26, 2005.

Accepted for publication December 29, 2005.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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