Figures
Abstract
Background
The success of ovarian follicle growth and ovulation is strictly related to the development of an adequate blood vessel network required to sustain the proliferative and endocrine functions of the follicular cells. Even if the Vascular Endothelial Growth Factor (VEGF) drives angiogenesis before ovulation, the local role exerted by Progesterone (P4) remains to be clarified, in particular when its concentration rapidly increases before ovulation.
Aim
This in vivo study was designed to clarify the effect promoted by a P4 receptor antagonist, RU486, on VEGF expression and follicular angiogenesis before ovulation, in particular, during the transition from pre to periovulatory follicles induced by human Chorionic Gonadotropins (hCG) administration.
Material and Methods
Preovulatory follicle growth and ovulation were pharmacologically induced in prepubertal gilts by combining equine Chorionic Gonadotropins (eCG) and hCG used in the presence or absence of RU486. The effects on VEGF expression were analyzed using biochemical and immunohistochemical studies, either on granulosa or on theca layers of follicles isolated few hours before ovulation. This angiogenic factor was also correlated to follicular morphology and to blood vessels architecture.
Results and Conclusions
VEGF production, blood vessel network and follicle remodeling were impaired by RU486 treatment, even if the cause-effect correlation remains to be clarified. The P4 antagonist strongly down-regulated theca VEGF expression, thus, preventing most of the angiogenic follicle response induced by hCG. RU486-treated follicles displayed a reduced vascular area, a lower rate of endothelial cell proliferation and a reduced recruitment of perivascular mural cells. These data provide important insights on the biological role of RU486 and, indirectly, on steroid hormones during periovulatory follicular phase. In addition, an in vivo model is proposed to evaluate how periovulatory follicular angiogenesis may affect the functionality of the corpus luteum (CL) and the success of pregnancy.
Citation: Mauro A, Martelli A, Berardinelli P, Russo V, Bernabò N, Di Giacinto O, et al. (2014) Effect of Antiprogesterone RU486 on VEGF Expression and Blood Vessel Remodeling on Ovarian Follicles before Ovulation. PLoS ONE 9(4): e95910. https://doi.org/10.1371/journal.pone.0095910
Editor: Anne Croy, Queen’s University, Canada
Received: December 16, 2013; Accepted: April 1, 2014; Published: April 22, 2014
Copyright: © 2014 Mauro et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by TERCAS Foundation 2009, Teramo, Italy. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Dominant preovulatory follicles are selected to grow from a pool of antral follicles during the ovarian cycle. This process leads to ovulation and CL formation [1]–[4]. Follicle selection success is strictly related to the development of a widespread blood vessel network required to sustain the enhanced proliferative and endocrine function of follicles [5]–[12]. Blood vessels allow growing follicles to acquire an increasing amount of nutrients, precursors, and hormones, as to release steroids and other regulating ovarian hormonal molecules to the systemic circulation [8], [13]. Several factors drive follicle angiogenesis indirectly controlling ovarian follicle development [6], [7], [11], [14]–[16]. VEGF is recognized to be a key molecule [13], [16], [17]. Indeed, its increased secretion, in addition to augmented vascular extension, is a necessary condition for large preantral follicles progression toward the antral stage [11], [18], [19]. Similarly, VEGF is up regulated in dominant follicle/s selection process leading to ovulation [7], [20]–[22]. Conversely, the process of follicle atresia is characterized both by VEGF and follicular blood vessel network reduction [14], [23]–[26].
VEGF, during gonadotropin surge, controls the crucial follicles transition from preovulatory to periovulatory stage that precedes ovulation [12], [16], [27]. In this time period of 24 or 44 h, depending on species, follicle profoundly changes its morphology and function [1], [25]. Luteinizing hormone (LH) surge induces a progressive disorganization of follicle basal membrane caused by proteolityc enzymes activation. Moreover, it modifies the steroid enzymatic pathway transforming an estrogen secreting preovulatory follicle to a P4 producing periovulatory follicle [28], [29]. In addition, follicular blood vessels undergo to dramatic changes. For the first time, large blood vessels appear in follicle walls and a higher blood flow is recorded [6], [7], [10], [27], [30]. Experimental evidences show that VEGF controls this vascular remodeling [10], [11], [21], [22], [27]. Indeed, its inhibition stops endothelial cell proliferation, impairs follicle angiogenesis preventing preovulatory follicle growth and ovulation [17], [22], [31], [32]. Although VEGF role on follicle development is clearly established, the mechanisms involved in its local expression remain to be clarified. Gonadotropins seem to link follicle growth and blood vessel remodeling through VEGF expression regulation [6], [7], [27], [33], [34]. However, it remains to be clarified whether gonadotropins directly affect VEGF expression or if this angiogenic factor is indirectly influenced by follicular steroidogenesis, as suggested by in vitro [33], [35] and in vivo experiments [36]. Amongst steroids, P4 may have a role in inducing VEGF expression [37], [38], endothelial cell proliferation [39]–[41], and angiogenesis [19], [42]. Indeed, the administration of P4 antagonist molecule, RU486, significantly decreased VEGF synthesis in vivo, in rat ovary [43] and in monkey endometrium [37]. Even if P4 influence on ovarian blood flow was demonstrated [44]–[47], during the periovulatory stage, when the LH surge induces a complete inversion of follicular steroid secretion [8], the role exerted by steroids on follicular VEGF synthesis still remains to be investigated.
To this aim, the present in vivo study was designed to study the effects induced by P4 antagonist administration, RU486 [48]–[53], on VEGF expression and angiogenesis during the transition from preovulatory to periovulatory follicles in gilts. This phase of follicle development was in vivo reproduced using a validated hormonal protocol [7], [27] able to promote antral follicular growth until the preovulatory stage (eCG injection) and ovulation (hCG treatment). Granulosa and theca VEGF contents were studied in selected follicles and its expression was correlated with follicle morphology and blood vessel organization assessed through histological, immunofluorescence and biochemical studies.
Materials and Methods
Ethical Committee
The experiments were approved by the Ethics Committee of the Universities of Teramo and Chieti-Pescara (Prot. 81/2011/CEISA/COM). All surgery was performed under sodium pentobarbital anesthesia, and all efforts were made to minimize suffering.
Animal Treatments and Ovary Collection
Follicular growth and ovulation were pharmacologically promoted into 20 prepubertal Large White gilts with a mean weight of 90.7±1.16 Kg (mean ± Standard Error, SE) using two sequentially intramuscular (i.m.) administrations of 1250 IU of eCG (Folligon; Intervet International B.V.-Boxmeer, Netherlands) and 750 IU of hCG (Corulon; Intervet International-Boxmeer, Netherlands), respectively [6], [7], [27]. The animals were divided in four experimental groups (5 animals each) as summarized in Fig. 1. Ovariectomies [27] were performed in order to collect preovulatory follicles 62 hours after eCG treatment (no hCG) and periovulatory ones collected 36 hours after hCG injection (Fig. 1B).
A) A schematic illustration of in vivo hormonal synchronization able to promote follicle growth and ovulation in prepubertal gilts. The periovulatory interval studied is showed in the box. B) Different experimental treatments and animal groups. All groups (n = 5 animals for each group) received eCG injection. Sixty-two hours after the eCG treatment, the first group (I°) was ovariectomized to obtain preovulatory follicles (no hCG), while the remaining groups received hCG treatment, with or without RU486. In detail, the second group (II°) received hCG injection dissolved in aqueous solution (*), as reagent control (CTR); the third group (III°) received hCG dissolved in corn oil (**), as experimental CTR, followed by administration of a corn oil alone 18 hours later; the fourth group (IV°) received hCG in combination with RU486 dissolved in corn oil and, 18 hours later, a second RU486 injection. Few hours before ovulation (36 hours after hCG treatments) the animals of these experimental groups were ovariectomized to obtain hCG, hCG+vehicle and hCG+RU486 periovulatory follicles. *commercial solvent **solvent for RU486.
In order to investigate the RU486 (Mifepristone, Sigma, St. Louis, USA) [48], [51] effects on periovulatory follicles, the hCG injection was carried out in combination with RU486. In particular, RU486 was solubilized in corn oil (10 ml) and i.m. injected at a concentration of 10 mg/kg [48]. RU486 half-life span is about 18–20 hours [51], [54], thus, two consecutive RU486 administrations were carried out: the first one in combination with hCG, the second one, 18 hours later, in corn oil (hCG+RU486) (Fig. 1B). The controls (CTR) were performed administrating hCG solubilized in aqueous solution (hCG, reagent CTR) or in corn oil (hCG+vehicle, experimental CTR) (Fig. 1B).
Sample Preparation
After ovariectomy, one ovary was fixed in 4% paraformaldehyde/phosphate-buffered saline (PBS; pH 7.4) for 12 h at 4°C, dehydrated and embedded in paraffin wax before performing histological and immunohistochemical analyses. The contra lateral ovary was collected to isolate single healthy preovulatory (8 mm<diameter ≥7 mm) and periovulatory follicles (11 mm ≤ diameter >8 mm), as previously reported [6], [27]. Each single follicle was opened under a stereomicroscope to collect follicular fluid (FF) and isolate follicular wall. Each follicular wall was dissected in order to separate granulosa cells and theca shell [6], [7], [27]. In detail, the follicle wall obtained from each structure was transferred into dissection medium and, with the aid of a small spatula, the granulosa layer was gently scraped away from the theca shell. The medium containing dispersed granulosa cells was collected and centrifuged, while the theca shell was vigorously vortexed and carefully washed in order to remove any possible granulosa cell contamination (as preliminary demonstrated by histological analysis) [6]. FF, granulosa and theca samples were individually stored in liquid nitrogen.
Morphological and Morphometric Analyses of the Follicular Response
Paraffin sections of 5 µm of thickness were serially collected on poly-L-lysine-coated slides before processing them for morphological, immunohistochemical (IHC) and morphometric investigations, according to Martelli et al. [11], [12], [27]. More in detail, at least three different sections/follicle were processed to analyze:
- tissue architecture, follicular diameter and oocyte’s health with Hematoxylin-Eosin (HE) staining;
- endothelial cells localization and vascular area (VA) extension using von Willenbrand Factor (vWF) IHC detection [27];
- VEGF distribution within granulosa and theca compartment using IHC detection [11];
- endothelial cell proliferation using vWF and Ki-67 double IHC (dIHC) detection [11], [27];
- blood vessel maturation and mural cell recruitment using vWF and alpha Smooth Muscle Actin (α-SMA) dIHC analysis [27]. The used antibodies are summarized in Table 1.
VA values were determined on the follicular wall (Tot VA) or on the inner and outer blood vessel network (iN and oN, respectively) by quantifying the relative vWF-positive area (µm2) determined within a defined field of 15000 µm2 [11], [12], [27]. The results were recorded on at least five different sections/follicle and expressed as mean values ± SE.
Endothelial cells proliferation index (PI) was expressed as the percentage of Ki67 positive endothelial cells (Ki-67-and vWF double-stained cells) on the total number of theca cells identified by 4′-6-diamidino-2-phenylindole dihydrochloride (DAPI) fluorescent nuclear staining [27].
Morphometric analyses were performed at x 400 Magnification according to Martelli et al. [11], [27] with an Axioscop-2plus-epifluorescence microscope (Zeiss) provided with an interactive and automatic image analyzer (Axiovision, Zeiss). Morphometrical analysis was then performed by using a KS300 computed image analysis system (Zeiss).
VEGF Biochemical Analysis
The levels and distribution of VEGF protein were analyzed in FF, granulosa and theca compartments.
VEGF Content in FF
VEGF content in FF samples was analyzed using a specific ELISA assay (Quantikine; R&D Systems, Minneapolis, MN) as previously described [6], [7], [27]. VEGF soluble levels recorded on at least five different samples of FF/hormonal treatment were expressed as ng/ml (mean values) ± SE.
VEGF Protein Content in Granulosa and Theca Compartments
Proteins extracted from granulosa or theca layers of each single isolated follicle (n = 5 follicles/gilt/treatment) were processed according to Martelli et al. [27]. Proteins (75 µg) were separated by 12% SDS-PAGE and electrophoretically transferred into a nitrocellulose membrane (Hybon C Extra; Amersham Pharmacia, Piscataway, NJ, USA) for Western Blot analysis [55]. Protein detection was performed by incubating the membranes with the primary polyclonal anti-human VEGF-antibody (2.5 µg/ml); Calbiochem, La Jolla, CA). The goat anti-rabbit immunoglobulins peroxidase-conjugated (IgG-HRP; 0.1 µg/ml, Santa Cruz Biotechnology Inc, Heidelberg, Germany) were finally used as secondary antibody. The membranes were re-probed with a monoclonal anti α-Tubulin (2 µg/ml, Sigma, St Louis, USA) and a goat anti-mouse IgG-HRP secondary antibody (0.1 µg/ml, Santa Cruz Biotechnology Inc, Heidelberg, Germany). The signals were detected with ECL Western Blot analysis system (Amersham Pharmacia, Amersham Pharmacia, Piscataway, NJ, USA). VEGF protein expression quantitative data were determined as the mean ratio of the optical density of specific bands normalized for the α-Tubulin expression through Advanced Image Data Analyzer (Rai Test; GMBH, Straubenhardt, Germany).
VEGF mRNA Content Recorded In Granulosa and Theca Layers Using Real Time PCR
Total RNA was extracted from granulosa and theca tissues of each single isolated follicle using TRI-Reagent (Sigma, S Louis, USA) as previously described [56]. Purified RNA was re-suspended in RNAse- free water and spectrophotometrically quantified (A260 nm). One µg of RNA was electrophoretically separated in 1% Agarose gel in order to determine RNA quality. The RNA was treated with DNase I digestion (Sigma) for 15 min at room temperature. One µg of total RNA was used for reverse transcription reaction with Oligo dT primer and BioScript™ (Bioline, London, UK).
Real-time quantitative PCR was performed with Stratagene Mx3000P instrument, (Stratagene, La Jolla, CA) using SYBR Green I dye detection accordingly to Barboni et al [56]. Detailed information on VEGF and β-Actin [57] gene primers, product length and cycles are reported in Table 2. The reaction components were prepared at a concentration of: 2.5 µl forward (0.25 µM) primer, 2.5 µl reverse (0.25 µM) primer (Table 2), 3.5 µl water and 12.5 µl Brilliant SYBR Green QPCR Master Mix 2X (Stratagene, La Jolla, CA). Three µl of cDNA were added to 22 µl of master mix. The real-time protocol used was: denaturation for 10 min at 95°C, 45 cycles at 95°C for 1 min, annealing at 60°C for 1 min, extension at 72°C for 1 min. Each sample/follicle was run in triplicate, and normalized to the housekeeping β-Actin gene mRNA level. The amplicons specificity was confirmed by dissociation curve and by 2% agarose gel electrophoresis. Each sample was run in triplicate and for quantitation of VEGF gene target the Comparative Ct Method was applied for all samples normalized to the control housekeeping β-Actin gene using the formula: 2−ΔΔC(t ) = 2 −(ΔCt control gene − ΔCt target gene). Statistical analysis was performed considering the last 3 experiments with a mean of 5 follicles/gilts/experiment.
Statistical Analysis
The data were expressed as mean ± SE of at least 3 independent experiments. The quantitative data obtained from the different hormonal treatments were, firstly assessed for normalcy of distribution, by D′Agostino and Pearson test. Then, the data sets were compared using ANOVA test followed, when necessary, by post-hoc Tukey test (GraphPad Prism 5, GraphPad Software, USA). The data were considered significant for p≤0.05.
Results
Ovarian Response to Hormonal Treatment
Macroscopic ovary evaluation showed that preovulatory and periovulatory follicular rate was not influenced by RU486 treatments as summarized in Table 3. In detail, preovulatory follicles (no hCG) were 21.15±2.17/animal and periovulatory ones were 19.85±2.21 (p>0.05), 21.02±1.74 (p>0.05) and 18.96±1.62 (p>0.05) in hCG, hCG+vehicle and hCG+RU486 animals, respectively.
HE staining (Fig. 2) confirmed that all isolated follicles were healthy and contained an oocyte surrounded by a cumulus cells continuous layer. They displayed a nucleus with a condensed chromatin and an uniform cytoplasm appearance (data non shown). Preovulatory follicles (n = 24; no hCG) showed a microscopic spherical regular architecture with a compact mural granulosa layer apposed on the basal membrane (Fig. 2A). Differently, periovulatory follicles isolated after hCG (data not shown) or hCG+vehicle (Fig, 2B) treatments displayed a typical dispersed granulosa layer, with infoldings of the theca projected towards the antrum. No vehicle effect was observed on periovulatory follicles. This periovulatory morphology was observed only in ∼ 30% of follicles isolated from RU486 treated gilts (n = 28; Fig, 2C, left image), while the remaining 70% maintained a structural organization that was similar to preovulatory follicles (Fig. 2C, right image).
Representative follicle images of tissue sections stained with HE. A) Preovulatory follicles (no hCG); B) hCG+vehicle periovulatory follicles; C) hCG+RU486 periovulatory follicles: left image, an example of a follicle isolated after RU486 treatment that displayed a typical periovulatory structures; right image, an example of a follicle isolated after RU486 treatment that showed a preovulatory-like morphology.
VEGF Protein Content-in Follicular Fluid and Granulosa/theca Compartments
FF, granulosa cells and theca layer were separately collected by each follicle with the aid of a stereomicroscope. As summarized in Table 4, VEGF content in FF was 15.96±0.53 ng/ml in preovulatory follicles (no hCG), resulted undetectable after both hCG treatments (hormone alone or solubilized in vehicle), and significantly lower in RU486 periovulatory follicles (0.98±0.07 ng/ml; Table 4). Granulosa VEGF protein expression, was similar in all analyzed preovulatory and periovulatory follicles (Fig. 3A). Instead, theca VEGF levels increased in both periovulatory hCG and hCG+vehicle follicles (Fig. 3B), while it did not change in hCG+RU486 follicles and it resulted similar to those recorded in preovulatory ones (Fig. 3B).
Representative Western Blot images of VEGF164 protein expression in A) granulosa cells and B) theca compartment of preovulatory follicles (no hCG), and hCG, hCG+vehicle, or hCG+RU486 periovulatory follicles. The histograms indicate VEGF164 densitometric values normalized for α-Tubulin. The values are expressed as mean ± SE of 3 independent experiments for a total of 15 follicles/treatment. a significantly different values (p<0.05) vs. no hCG treatment * significantly different values (p<0.05) vs. hCG+vehicle treatment.
VEGF Protein Distribution in Follicular Compartments
VEGF protein distribution within granulosa and theca layers was investigated using IHC, as shown in Fig. 4. In all categories of follicles, VEGF was present within several granulosa cells or accumulated amongst them. By contrast, VEGF localization in theca compartment resulted influenced by hormonal treatments. More in detail, in preovulatory follicles (no hCG, Fig. 4A) VEGF was exclusively localized near the blood vessels. Differently, both periovulatory hCG (data not shown) and hCG+vehicle follicles (Fig. 4B) showed a preferential extracellular matrix VEGF distribution close to the basal membrane and to blood vessels. A similar theca VEGF distribution was observed in RU486 periovulatory follicles (Fig. 4C), even if in those follicles that maintained a preovulatory-like organization its distribution close to the blood vessels appeared lower, if not absent (Fig. 4C, right image).
Representative follicle images displaying VEGF protein localization detected by IHC. VEGF was revealed by black positivity (DAB signal) detected within cells (black arrows) or deposited amongst the extracellular matrix. A) preovulatory follicle (no hCG), B) hCG+vehicle periovulatory follicle, and C) hCG+RU486 follicle: left image, an example of RU486 follicle that displayed a typical periovulatory structure; right image, an example of RU486 follicle that showed a preovulatory-like morphology.
VEGF mRNA Expression in Granulosa and Theca Layers
As shown in Fig. 5, VEGF mRNA expression significantly decreased in granulosa cells after hCG injection, regardless of the treatments used (Fig. 5A). Differently, VEGF mRNA levels increased in theca layer after hCG injection alone or with vehicle (Fig. 5B). By contrast, VEGF up regulation was absent in theca layers of follicles obtained from RU486 treated animals (Fig. 5B, hCG+RU486 vs hCG+vehicle, p≤0.05) and its level remained similar to those of preovulatory ones (Fig. 5B, hCG+RU486 vs. no hCG, p≥0.05).
Representative images displaying VEGF164 mRNA expression analyzed by Real Time PCR. A) granulosa and B) theca compartment of preovulatory follicles (no hCG) and hCG, hCG+vehicle, or hCG+RU486 periovulatory follicles. The histogram indicates VEGF values quantified by a Comparative Ct method and expressed as the mean ± SE of 3 independent experiments for a total of 15 follicles/treatment. Amplicons specificity was confirmed by 2% agarose gel electrophoresis. asignificantly different values (p<0.05) vs. no hCG treatment. *significantly different values (p<0.05) vs. hCG+vehicle treatment.
Blood Vessel Remodeling during RU486 Treatment
The effects of RU486 on blood vessel organization (Fig. 6), VA and endothelial cell proliferation (Fig. 7) were evaluated using IHC. Preovulatory follicles (no hCG) displayed a blood vessel network organized into two concentric structures connected to each other by anastomotic vessels. The iN VA (2661.82±89.46 µm2/field, Fig. 7A), characterized by capillaries close to the basal membrane (Fig. 6), was lower than the oN VA (3751.07±89.44 µm2/field, Fig. 7A). The Tot VA significantly increased after both hCG and hCG+vehicle treatments (Fig. 7A, p≤0.01). In particular, the oN VA dramatically increased (Fig. 7A) becoming ∼70% of Tot VA for the presence of new large blood vessels (Fig. 6).
Representative images of double IHC performed on preovulatory (no hCG) and hCG+vehicle or hCG+RU486 periovulatory follicles to reveal the endothelial marker vWF (red fluorescence, Cy3) and, the smooth muscle actin (α-SMA) antigen (green fluorescence, Alexa Fluor 488). Cell nuclei were counterstained with DAPI (blue fluorescence) to easily distinguish granulosa (G), iN, and oN blood vessel networks localized in theca compartment. In hCG+RU486 image it is shown an example of the α-SMA and vWF distribution in a preovulatory-like organized follicle (“pre-like”), while in the insert panels RU486 examples are shown with the typical periovulatory morphology (“peri” in the left upper corner).
A) The histogram shows the VA values calculated, in preovulatory follicles (no hCG), and in hCG, hCG+vehicle or hCG+RU486 periovulatory follicles, considering the vWF-positive area/15000 µm2 of theca field. The VAs were divided in iN (open bars) and oN (shaded bars), on the basis of their localization. The total VA (solid bars) expressed the iN VA+oN VA. The preovulatory–like (“pre-like”) and periovulatory (“peri”) hCG+RU486 follicle VAs were calculated separately. The data expressed as mean values ± SE were obtained from 15 different follicles/treatment. astatistically different values (p<0.05) of each VA vs. no hCG preovulatory follicles. B) The histogram shows the endothelial cell PI given, after dIHC analyses, by the percentage of Ki 67 (a cell proliferation marker) positive cells that co-localize with the endothelial cells (anti-vWF marker) recorded in a standardized field (15.000 µm2 of theca layer field). Total PI (solid bars) was divided in iN (open bars) and oN (shaded bars) PIs, on the basis of their localization. The data expressed as mean values ± SE were obtained from 15 different follicles/treatment. astatistically different values (p<0.01) of each PI vs. no hCG preovulatory follicles.
Total and oN VAs similarly increased in follicles that acquired after RU486 treatment a periovulatory organization (Fig. 6 insert image and Fig. 7A, “peri”). On the contrary, VAs did not change in those follicles that maintained a preovulatory-like architecture which showed few large vessels within the theca (Fig. 6 and Fig. 7A, “pre-like”).
The vWF (red stain) and α-SMA (green stain) double-immunolabeling, performed to describe blood vessel maturity and perivascular recruited cells, showed a clear RU486 treatment effect. (no hCG) Blood vessels preovulatory follicles (iN and oN) did not display α-SMA immunopositivity (Fig. 6). Instead, perivascular α-SMA positive cells appeared in iN and oN of periovulatory follicles obtained after hCG (data not shown) and hCG+vehicle treatments (Fig. 6). In particular, the iN displayed an α-SMA immunopositivity close to the endothelial cells (vWF positive cells) forming a “chain-like” organization. By contrast, α-SMA positivity was observed at the periphery of the large blood vessel walls localized in oN (Fig. 6).
A similar distribution of α-SMA- positive cells was observed in those follicles isolated by RU486 treated animals that acquired a periovulatory organization (Fig. 6 insert image, “peri”). By contrast, the follicles maintaining the preovulatory-like architecture (Fig. 6, “pre-like”) did not display α-SMA positivity into the iN, and showed a reduced α-SMA perivascular immunoreaction in the oN large blood vessels. Rarely, these large blood vessels displayed a continuous α-SMA perivascular layer (Fig. 6, “pre-like”).
vWF and Ki-67 Staining
The vWF and Ki-67 double immunolabeling was used to quantify the proliferating endothelial cells in the different classes of follicles (Fig. 7B). Preovulatory (no hCG) and periovulatory follicles (hCG and hCG+vehicle) displayed endothelial Ki-67 positive cells in the iN and oN. The incidence of proliferating endothelial cells significantly decreased only in follicles isolated from RU486 treated animals that maintaining the preovulatory-like architecture (Fig. 7B, “pre-like”).
Discussion
The present study was performed to clarify the role of RU486 on VEGF-dependent ovarian angiogenesis that occurs in vivo in dominant follicles after hCG administration (to mimic LH surge). Our results demonstrate, for the first time, that VEGF production, blood vessel network, and follicle tissue remodeling were all affected by the in vivo administration of RU486 during the phase of transition from pre to periovulatory follicle, characterized by increasing levels of P4 [28], [29], even if the cause-effect correlation between these events remains to be clarified. In addition to pregnancy [52], [53], [58], [59], ovulation [60]–[65], and sperm-oocyte recognition [66], a new RU486 negative effect was demonstrated on reproductive events. Indeed, RU486 administration was able to impair periovulatory follicle development and angiogenesis induced by hCG. Its effect appeared to be compartment and vascular (oN and/or iN) network-dependent. In particular, RU486 treatment down-regulated VEGF expression within the theca compartment, while, it did not affect granulosa and FF VEGF content. Independently of RU486 injection, FF VEGF levels dramatically decreased after hCG treatment. This result could be apparently contradictory, since low FF VEGF levels were recorded both in RU486 and hGC periovulatory follicles characterized by low and high theca VEGF expression respectively. This observation, may be explained hypothesizing that hCG treatment, induces the synthesis of VEGF low bioavailability isoforms in follicles. This suggestion is in agreement with the expression of high molecular weight (i.e. VEGF189) VEGF isoforms with a lower solubility evidence in endometrium stimulated by a modified steroid milieu [67]. Alternatively, VEGF bioavailability could be affected by local changes of heparin or cell surface heparan sulphate concentration, as previously supposed by Robinson & Stringer [68]. Regardless the mechanism, RU486 was able to selectively inhibit VEGF expression, triggered by hCG, within theca compartment of periovulatory follicles. This inhibitory effect may be a consequence of the antiprogesterone action of RU486, as reported in bovine [69], human [70] and mink [71], and in another reproductive tissue such as endometrium [37], [38], [42], [72] exposed to low concentrations of P4. As a consequence, the RU486 follicles displayed an incomplete development of the blood vessel network. It is interesting to note as RU486 treatment did not influence the total VA but, specifically, affected oN VA that displayed a reduced extension. This important inhibitory RU486 influence was exclusively observed in 70% of the follicles that did not acquire the typical periovulatory organization. These results confirm that morphological and vascular remodeling recognize controlling cross-talk mechanisms [73]–[78]. On the contrary, RU486 treatment did not affect iN VA that probably remained regularly controlled by the unchanged FF and granulosa compartment VEGF levels. The presence of iN VA may have an essential functional role since it could maintain an adequate trophic supply of the germinal structure. Indeed, its persistence is vital to ensure the correct oxygen, precursors and metabolites supply either to the avascular granulosa compartment or to the oocyte [1], [6], [7], [27].
Though, the RU486 inhibitory effects described on endothelial cell proliferation and mural cell recruitment can not be exclusively explained with the theca VEGF down-regulation. Indeed, RU486 treatment induced these responses either the oN or iN. The inhibition of endothelial cell proliferation could be a consequence either of VEGF lower levels, accordingly to other Authors [75], [76], as of RU486 antagonist role on P4 receptor. This hypothesis seems to be supported by the observations of P4 receptors expression in Huvec cells [79], [80] and by the P4-dependent inhibition of endothelial cell proliferation, already demonstrated in endometrial blood vessels in mouse [42] and in uterine vasculature in pig [38]. RU486 inhibitory role on mural cell recruitment affected also the consequent blood vessel maturity. Human CG injection in periovulatory follicles promotes the maturation of blood vessels by recruiting mural cells (α–SMA positive cells). These cells in the oN surround large blood vessels, while in iN capillary network acquire the typical “chain-like organization” [27]. The RU486 negative influence on blood vessel maturity could be related to a direct effect of P4, as already suggested in mouse [42], or as a consequence of VEGF lower expression. It is known that this angiogenic factor indirectly modulates blood vessel maturation in other systems by influencing the secretion of key factors such as Angiopoietins/Tie-2 receptor [74], [75], [81], [82], platelet-derived growth factor (PDGF)-B/PDGF receptor beta [76], and the transforming growth factor beta (TGF beta) [76]. Regardless the mechanisms involved, blood vessels immaturity after RU486 treatment may impair the physiological transition from periovulatory follicle to CL structure. The presence of smooth muscle in follicle blood vessels is essential for the control exerted by nervous system or local hormones on the ovarian function [27]. In particular, the large vessels with a smooth muscle mural wall absence in the oN could probably impair blood flow required for ovulation and the subsequent CL development [83], [84]. Moreover, α–SMA positive cells absence within the capillaries of the iN could negatively affect blood vessel stabilization and haemodynamic processes necessary to support the early stage of CL growth [85]. In this context, further experiments will be required to demonstrate if and how the low maturity of follicular blood vessels could influence ovulation and CL function.
In conclusion, the obtained results in the present work demonstrate RU486 negative influences on VEGF expression, vascular and tissue remodeling, providing important insights on the biological in vivo role of this P4 antagonist during the transition from the preovulatory to the periovulatory stage. Even if the cause-effect correlation amongst angiogenesis and impaired follicle development has not been demonstrated, these results propose an in vivo model that allows to study the effect of an incomplete follicle maturation on female reproduction success.
Acknowledgments
We thank Delia Nardinocchi, Domenica Cocciolone and Maura Turriani for their valuable technical assistance.
Author Contributions
Conceived and designed the experiments: A. Mauro A. Martelli BB MM. Performed the experiments: A. Mauro A. Martelli PB VR NB OD. Analyzed the data: A. Mauro A. Martelli BB PB VR MM. Contributed reagents/materials/analysis tools: BB MM. Wrote the paper: A. Mauro A. Martelli BB. Critically revised paper: BB MM.
References
- 1. Moor RM, Seamark RF (1986) Cell signaling, permeability, and microvasculatory changes during antral follicle development in mammals. J Dairy Sci 69: 927–943.
- 2. Zeleznik AJ (2001) Follicle selection in primates: “many are called but few are chosen”. Biol Reprod 65: 655–659.
- 3. Adams GP, Singh J, Baerwald AR (2012) Large animal models for the study of ovarian follicular dynamics in women. Theriogenology 78(8): 1733–48.
- 4. Baerwald AR, Adams GP, Pierson RA (2012) Ovarian antral folliculogenesis during the human menstrual cycle: a review. Hum Reprod Update 18: 73–91.
- 5. Risau W (1997) Mechanisms of angiogenesis. Nature 386: 671–674.
- 6. Barboni B, Turriani M, Galeati G, Spinaci M, Bacci ML, et al. (2000) Vascular endothelial growth factor production in growing pig antral follicles. Biol Reprod 63: 858–864.
- 7. Mattioli M, Barboni B, Turriani M, Galeati G, Zannoni A, et al. (2001) Follicle activation involves vascular endothelial growth factor production and increased blood vessel extension. Biol Reprod 65: 1014–1019.
- 8.
Geva E, Jaffe RB (2004) Ovarian angiogenesis In: Leung P, Adashi E, eds. The Ovary. 2nd ed. San Diego (CA): Elsevier Science.
- 9.
Gougeon A (2004) Dynamics for human follicular growth: morphologic, dynamic and functional aspects. In: Leung P, Adashi E, eds. The Ovary. 2nd ed. San Diego (CA): Elsevier Academic Press 25–43.
- 10. Hunter MG, Robinson RS, Mann GE, Webb R (2004) Endocrine and paracrine control of follicular development and ovulation rate in farm species. Anim Reprod Sci 82–83: 461–477.
- 11. Martelli A, Bernabo N, Berardinelli P, Russo V, Rinaldi C, et al. (2009) Vascular supply as a discriminating factor for pig preantral follicle selection. Reproduction 137(1): 45–58.
- 12. Martelli A, Palmerini MG, Russo V, Rinaldi C, Bernabo N, et al. (2009) Blood vessel remodeling in pig ovarian follicles during the periovulatory period: an immunohistochemistry and SEM-corrosion casting study. Reprod Biol Endocrinol 7: 72.
- 13. Geva E, Jaffe RB (2000) Role of vascular endothelial growth factor in ovarian physiology and pathology. Fertil Steril 74: 429–438.
- 14. Plendl J (2000) Angiogenesis and vascular regression in the ovary. Anat Histol Embryol 29: 257–266.
- 15. Shimizu T, Kawahara M, Abe Y, Yokoo M, Sasada H, et al. (2003) Follicular microvasculature and angiogenic factors in the ovaries of domestic animals. J Reprod Dev 49(3): 181–192.
- 16. Robinson RS, Woad KJ, Hammond AJ, Laird M, Hunter MG, et al. (2009) Angiogenesis and vascular function in the ovary. Reproduction 138: 869–881.
- 17. Fraser HM (2006) Regulation of the ovarian follicular vasculature. Reprod Biol Endocrinol 4: 18.
- 18. Danforth DR, Arbogast LK, Ghosh S, Dickerman A, Rofagha R, et al. (2003) Vascular endothelial growth factor stimulates preantral follicle growth in the rat ovary. Biol Reprod 68: 1736–1741.
- 19. Kaczmarek MM, Schams D, Ziecik AJ (2005) Role of vascular endothelial growth factor in ovarian physiology – an overview. Reprod Biol 5(2): 111–136.
- 20. Stouffer RL, Martinez-Chequer JC, Molskness TA, Xu F, Hazzard TM (2001) Regulation and action of angiogenic factors in the primate ovary. Arch Med Res 32: 567–575.
- 21. Wulff C, Wiegand SJ, Saunders PT, Scobie GA, Fraser HM (2001) Angiogenesis during follicular development in the primate and its inhibition by treatment with truncated Flt-1-Fc (vascular endothelial growth factor Trap(A40)). Endocrinology 142: 3244–3254.
- 22. Wulff C, Wilson H, Wiegand SJ, Rudge JS, Fraser HM (2002) Prevention of thecal angiogenesis, antral follicular growth, and ovulation in the primate by treatment with vascular endothelial growth factor Trap R1R2. Endocrinology 143: 2797–2807.
- 23. Greenwald GS (1989) Temporal and Topographic Changes in DNA Synthesis after Induced Follicular Atresia. Biol Reprod 40: 175–181.
- 24. Jablonka-Shariff A, Fricke PM, Grazul-Bilska AT, Reynolds LP, Redmer DA (1994) Size, Number, Cellular Proliferation, and Atresia of Gonadotropin-lnduced Follicles in Ewes. Biol Reprod 51: 531–540.
- 25. Redmer DA, Reynolds LP (1996) Angiogenesis in the ovary. Rev Reprod 1: 182–192.
- 26. Berardinelli P, Russo V, Martelli A, Nardinocchi D, Di Giacinto O, et al. (2004) Colocalization of DNA fragmentation and caspase-3 activation during atresia in pig antral follicles. Anat Histol Embryol 33(1): 23–7.
- 27. Martelli A, Berardinelli P, Russo V, Mauro A, Bernabò N, et al. (2006) Spatio-temporal analysis of vascular endothelial growth factor expression and blood vessel remodelling in pig ovarian follicles during the periovulatory period. J Mol Endocrinol 36(1): 107–19.
- 28. Berisha B, Steffl M, Welter H, Kliem H, Meyer HH, et al. (2008) Effect of the luteinising hormone surge on regulation of vascular endothelial growth factor and extracellular matrix-degrading proteinases and their inhibitors in bovine follicles. Reprod Fertil Dev 20(2): 258–68.
- 29. Fortune JE, Willis EL, Bridges PJ, Yang CS (2009) The periovulatory period in cattle: progesterone, prostaglandins, oxytocin and ADAMTS proteases. Anim Reprod 6(1): 60–71.
- 30. Acosta TJ, Hayashi KG, Ohtani M, Miyamoto A (2003) Local changes in blood flow within the preovulatory follicle wall and early corpus luteum in cows. Reproduction 125: 759–767.
- 31. Zimmermann RC, Xiao E, Bohlen P, Ferin M 2002 Administration of antivascular endothelial growth factor receptor 2 antibody in the early follicular phase delays follicular selection and development in the rhesus monkey. Endocrinology 143: 2496–2502.
- 32. Fraser HM, Duncan WC 2009 Regulation and manipulation of angiogenesis in the ovary and endometrium. Reprod Fertil Dev 21: 377–392.
- 33. Hazzard TM, Molskness TA, Chaffin CL, Stouffer RL (1999) Vascular endothelial growth factor (VEGF) and angiopoietin regulation by gonadotrophin and steroids in macaque granulosa cells during the peri-ovulatory interval. Mol Hum Reprod 5: 1115–1121.
- 34. Gutman G, Barak V, Maslovitz S, Amit A, Lessing JB, et al. (2008) Regulation of vascular endothelial growth factor-A and its soluble receptor sFlt-1 by luteinizing hormone in vivo: implication for ovarian follicle angiogenesis. Fertil Steril 89: 922–926.
- 35. Shimizu T, Miyamoto A (2007) Progesterone induces the expression of vascular endothelial growth factor (VEGF) 120 and Flk-1, its receptor, in bovine granulosa cells. Anim Reprod Sci 102: 228–237.
- 36. Hyder SM, Stancel GM (1999) Regulation of Angiogenic Growth Factors in the Female Reproductive Tract by Estrogens and Progestins. Mol Endocrinol 13(6): 806–811.
- 37. Greb RR, Heikinheimo O, Williams RF, Hodgen GD, Goodman AL (1997) Vascular endothelial growth factor in primate endometrium is regulated by oestrogen-receptor and progesterone-receptor ligands in vivo. Hum Reprod 12: 1280–1292.
- 38. Bailey DW, Dunlap KA, Frank JW, Erikson DW, White BG, et al. (2010) Effects of long-term progesterone on developmental and functional aspects of porcine uterine epithelia and vasculature: progesterone alone does not support development of uterine glands comparable to that of pregnancy. Reproduction 140: 583–594.
- 39. Vazquez F, Rodriguez-Manzaneque JC, Lydon JP, Edwards DP, O’Malley BW, et al. (1999) Progesterone regulates proliferation of endothelial cells. J Biol Chem 274: 2185–2192.
- 40. Hsu SP, Ho PY, Juan SH, Liang YC, Lee WS (2008) Progesterone inhibits human endothelial cell proliferation through a p53-dependent pathway. Cell Mol Life Sci 65: 3839–3850.
- 41. Hsu SP, Lee WS (2011) Progesterone receptor activation of extranuclear signaling pathways in regulating p53 expression in vascular endothelial cells. Mol Endocrinol 25(3): 421–32
- 42. Girling JE, Lederman FL, Walter LM, Rogers PA (2007) Progesterone, but not estrogen, stimulates vessel maturation in the mouse endometrium. Endocrinology 148: 5433–5441.
- 43. Ishikawa K, Ohba T, Tanaka N, Iqbal M, Okamura Y, et al. (2003) Organ-specific production control of vascular endothelial growth factor in ovarian hyperstimulation syndrome-model rats. Endocr J 50: 515–525.
- 44. Ricke WA, Redmer DA, Reynolds LP (1999) Growth and cellular proliferation of pig corpora lutea throughout the oestrous cycle. J Reprod Fertil 117: 369–377.
- 45. Fraser HM, Lunn SF (2001) Regulation and manipulation of angiogenesis in the primate corpus luteum. Reproduction 121: 355–362.
- 46. Luttgenau J, Beindorff N, Ulbrich SE, Kastelic JP, Bollwein H (2011) Low plasma progesterone concentrations are accompanied by reduced luteal blood flow and increased size of the dominant follicle in dairy cows. Theriogenology 76: 12–22.
- 47. Pancarci SM, Gungor O, Atakisi O, Cigremis Y, Ari UC, et al. (2011) Changes in follicular blood flow and nitric oxide levels in follicular fluid during follicular deviation in cows. Anim Reprod Sci 123: 149–156.
- 48. Baulieu EE (1989) Contragestion and other clinical applications of RU 486, an antiprogesterone at the receptor. Science 245: 1351–1357.
- 49. Spitz IM, Croxatto HB, Lahteenmaki P, Heikinheimo O, Bardin CW (1994) Effect of Mifepristone on Inhibition of Ovulation and Induction of Luteolysis. Hum Reprod 9: 69–76.
- 50. Cadepond F, Ulmann A, Baulieu EE (1997) RU486 (mifepristone): Mechanisms of action and clinical uses. Annu Rev Med 48: 129–156.
- 51. Heikinheimo O, Kekkonen R, Lahteenmaki P (2003) The pharmacokinetics of mifepristone in humans reveal insights into differential mechanisms of antiprogestin action. Contraception 68: 421–426.
- 52.
Mathew D, Sellner E, Okamura C, Geisert R, Anderson L, et al.. (2009) Effect of progesterone antagonist RU486 on uterine progesterone receptor mRNA expression, embryonic development and ovarian function during early pregnancy in pigs. Soc Reprod Fertil Suppl 66: 333–334.
- 53. Mathew DJ, Sellner EM, Green JC, Okamura CS, Anderson LL, et al. (2011) Uterine Progesterone Receptor Expression, Conceptus Development, and Ovarian Function in Pigs Treated with RU 486 During Early Pregnancy. Biol Reprod 84: 130–139.
- 54. Sarkar NN (2002) Mifepristone: bioavailability, pharmacokinetics and use-effectiveness. Eur J Obstet Gynecol Reprod Biol 101: 113–120.
- 55. Towbin H, Staehelin T, Gordon J (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc Natl Acad Sci USA 76(9): 4350–4.
- 56. Barboni B, Mangano C, Valbonetti L, Marruchella G, Berardinelli P, et al. (2013) Synthetic Bone Substitute Engineered with Amniotic Epithelial Cells Enhances Bone Regeneration after Maxillary Sinus Augmentation. PLoS One 8(5): e63256
- 57. Shimizu T, Jiang JY, Sasada H, Sato E (2002) Changes of messenger RNA expression of angiogenic factors and related receptors during follicular development in gilts. Biol Reprod 67(6): 1846–1852.
- 58. Bouchard P, Chabbert-Buffet N, Fauser BC (2011) Selective progesterone receptor modulators in reproductive medicine: pharmacology, clinical efficacy and safety. Fertil Steril 96: 1175–1189.
- 59. Liang H, Gao ES, Chen AM, Luo L, Cheng YM, et al. (2011) Mifepristone-induced abortion and vaginal bleeding in subsequent pregnancy. Contraception 84: 609–614.
- 60. Abe T, Toida D, Satoh H, Yonezawa T, Kawaminami M, et al. (2011) An early single dose of progesterone agonist attenuates endogenous progesterone surge and reduces ovulation rate in immature rat model of induced ovulation. Steroids 76(10–11): 1116–25
- 61. Kawano T, Okamura H, Tajima C, Fukuma K, Katabuchi H (1988) Effect of RU 486 on luteal function in the early pregnant rat. J Reprod Fertil 83: 279–285.
- 62. Tanaka N, Iwamasa J, Matsuura K, Okamura H (1993) Effects of progesterone and anti-progesterone RU486 on ovarian 3 beta-hydroxysteroid dehydrogenase activity during ovulation in the gonadotrophin-primed immature rat. J Reprod Fertil 97: 167–172.
- 63. Brown A, Cheng L, Lin S, Baird DT (2002) Daily low-dose mifepristone has contraceptive potential by suppressing ovulation and menstruation: a double-blind randomized control trial of 2 and 5 mg per day for 120 days. J Clin Endocrinol Metab 87: 63–70.
- 64. Brown A, Williams A, Cameron S, Morrow S, Baird DT (2003) A single dose of mifepristone (200 mg) in the immediate preovulatory phase offers contraceptive potential without cycle disruption. Contraception 68: 203–209.
- 65. Weisberg E, Croxatto HB, Findlay JK, Burger HG, Fraser IS (2011) A randomized study of the effect of mifepristone alone or in conjunction with ethinyl estradiol on ovarian function in women using the etonogestrel-releasing subdermal implant, Implanon(R). Contraception 84: 600–608.
- 66. Siqueira LC, Barreta MH, Gasperin B, Bohrer R, Santos JT, et al. (2012) Angiotensin II, progesterone, and prostaglandins are sequential steps in the pathway to bovine oocyte nuclear maturation. Theriogenology 77: 1779–1787.
- 67. Ancelin M, Buteau-Lozano H, Meduri G, Osborne-Pellegrin M, Sordello S, et al. (2002) A dynamic shift of VEGF isoforms with a transient and selective progesterone-induced expresion of VEGF 189 regulates angiogenesis and vascular permeability in human uterus. PNAS 99: 6023–6028.
- 68. Robinson CJ, Stringer SE (2001) The splice variants of vascular endothelial growth factor (VEGF) and their receptors. J Cell Sci 114: 853–865.
- 69. Sagsoz H, Saruhan BG (2011) The expression of vascular endothelial growth factor and its receptors (flt1/fms, flk1/KDR, flt4) and vascular endothelial growth inhibitor in the bovine uterus during the sexual cycle and their correlation with serum sex steroids. Theriogenology 75: 1720–1734.
- 70. Sugino N, Kashida S, Karube-Harada A, Takiguchi S, Kato H (2002) Expression of vascular endothelial growth factor (VEGF) and its receptors in human endometrium throughout the menstrual cycle and in early pregnancy. Reproduction 123: 379–387.
- 71. Lopes FL, Desmarais J, Gevry NY, Ledoux S, Murphy BD (2003) Expression of vascular endothelial growth factor isoforms and receptors Flt-1 and KDR during the peri-implantation period in the mink, Mustela vison. Biol Reprod 68: 1926–1933.
- 72. Kaczmarek MM, Waclawik A, Blitek A, Kowalczyk AE, Schams D, et al. (2008) Expression of the vascular endothelial growth factor-receptor system in the porcine endometrium throughout the estrous cycle and early pregnancy. Mol Reprod Dev 75: 362–372.
- 73. Gottsch ML, Van Kirk EA, Murdoch WJ (2002) Role of matrix metalloproteinase 2 in theovulatory folliculo-luteal transition of ewes. Reproduction 124(3): 347–52.
- 74. Armulik A, Abramsson A, Betsholtz C (2005) Endothelial/pericyte interactions. Circ Res 97: 512–523.
- 75. Girling JE, Rogers PA (2009) Regulation of endometrial vascular remodelling: role of the vascular endothelial growth factor family and the angiopoietin-TIE signalling system. Reproduction 138: 883–893.
- 76. Jain RK (2003) Molecular regulation of vessel maturation. Nat Med 9: 685–693.
- 77. Ferrara N, Davis-Smyth T (1997) The biology of vascular endothelial growth factor. Endocr Rev 18: 4–25.
- 78. Ferrara N (2010) Binding to the extracellular matrix and proteolytic processing: two key mechanisms regulating vascular endothelial growth factor action. Mol Biol Cell 21: 687–690.
- 79. Toth B, Saadat G, Geller A, Scholz C, Schulze S, et al. (2008) Human umbilical vascular endothelial cells express estrogen receptor beta (ERbeta) and progesterone receptor A (PR-A), but not ERalpha and PR-B. Histochem Cell Biol 130: 399–405.
- 80. Oviedo PJ, Sobrino A, Novella S, Rius C, Laguna-Fernandez A, et al. (2011) Progestogens reduce thromboxane production by cultured human endothelial cells. Climacteric 14(1): 41–48.
- 81. Singh H, Milner CS, Aguilar Hernandez MM, Patel N, Brindle NP (2009) Vascular endothelial growth factor activates the Tie family of receptor tyrosine kinases. Cell Signal 21: 1346–1350.
- 82. Herbert SP, Stainier DY (2011) Molecular control of endothelial cell behaviour during blood vessel morphogenesis. Nat Rev Mol Cell Biol 12(9): 551–64.
- 83. Bergers G, Song S (2005) The role of pericytes in blood-vessel formation and maintenance. Neuro Oncol 7: 452–464.
- 84.
Betsholtz C, Lindblom P, Gerhardt H (2005) Role of pericytes in vascular morphogenesis. EXS 115–125.
- 85.
Murphy BD, Gevry N, Ruiz-Cortes T, Cote F, Downey BR, et al.. (2001) Formation and early development of the corpus luteum in pigs. Reprod Suppl 58: 47–63.