| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Gamete Biology |
Unité de Physiologie de la Reproduction et des Comportements, Institut National de la Recherche Agronomique, 37380 Nouzilly, France
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
cumulus cells, gamete biology, growth hormone, oocyte development, ovum
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
GH may act directly on the ovary. In perfused rabbit ovary, GH stimulated follicle growth, oocyte maturation, and ovarian estradiol production [3]. In vitro, GH stimulates granulosa cell proliferation and steroidogenesis [4–9]. The effects of GH on steroidogenesis depends on follicular developmental stage [6, 9]. Furthermore, the GH receptor was localized in follicle cells in most species (rat [10], bovine [11, 12], ovine [13], porcine [14], human [15]).
In vitro, the effects of GH on oocyte maturation (IVM) were mainly studied in the bovine. Izadyar et al. [16] reported an acceleration of IVM of cumulus-enclosed oocytes, the induction of cumulus expansion, and the promotion of subsequent embryonic development. They also demonstrated enhanced migration of cortical granules and sperm aster formation [17]. Iga et al. [18] showed increased rates of nuclear maturation and blastocyst formation of bovine oocytes in the presence of GH, but they did not observe GH-induced cumulus expansion nor increased rates of fertilization. Nevertheless, it was shown recently that the improvement of bovine oocyte maturation and fertilization rates by GH was dependent on the culture medium [19]. This could account for the absence of positive effect of the GH on bovine oocyte maturation observed by Sirotkin [7]. Hagen and Graboski [20] suggested that GH accelerated the maturation of porcine oocytes in vitro because the percentage of oocytes activated in an electroporation chamber after 24 h of IVM was increased in the presence of GH. An accelerated meiotic maturation was also observed in rat [21] and in blue fox [22] oocytes when GH was added to the maturation medium. However, GH had no effect on canine oocyte maturation [23] and felid oocyte maturation, fertilization, and developmental potential [24]. Nevertheless, the high concentration of GH used in this study could be responsible for the negative results.
Scarce studies have been performed in species other than bovine. However, GH may improve in vitro embryo production (IVP) results in domestic animals such as equine and porcine. Indeed, major problems are encountered with IVP in these two species. The results of in vitro maturation of equine oocytes are still low in comparison with other mammalian species. Numerous studies have been conducted to improve the rate of maturation of equine oocytes, and positive effects of LH, FSH, estradiol, and epidermal growth factor (EGF) have been described [25, 26]. However, the low rates of oocyte maturation remain limiting for the development of in vitro fertilization (IVF) and embryo development (IVD) techniques. Improving the conditions of equine IVM are needed for further steps of embryo production in vitro. In porcine oocytes, the use of growth factors and gonadotropins improved nuclear and cytoplasmic maturation. Nevertheless, polyspermy is still a major obstacle, suggesting incomplete cytoplasmic maturation [27 for review].
The way of action of GH is not well known. GH receptor is expressed in cumulus cells and in oocyte in bovine and rat [10–12, 28], but the effects of GH on oocyte maturation are exerted via cumulus cells in the two species [21, 12]. In the rat, GH effects are mediated by IGF-I [21]. In bovine, some studies showed that GH acts via IGF-I [29], while others showed that GH acts via a cAMP-mediated pathway without mediation of IGF-I [12, 30]. To our knowledge, no studies have been published in other species. Moreover, the mechanism of GH action on cumulus expansion is not known.
In the process of cumulus expansion, cumulus cells secrete hyaluronan that accumulates among the cells. Recently, three kinds of mammalian genes encoding hyaluronan synthases (Has 1, Has 2, and Has 3) have been identified [31–36]. Several mammalian cell lines transfected with their cDNAs exhibited a marked increase of hyaluronan production, indicating an important role for these enzymes in hyaluronan biosynthesis [32, 34, 35, 37]. Moreover, the expression of hyaluronan synthases has been reported in mouse and porcine cumulus [36, 38]. The factors modulating the Has expression have hardly been investigated. The cumulus expansion is also accompanied by modifications of gap junctions, which contain transmembrane channels formed by hexamers of proteins belonging to the connexin family. Bovine and mouse cumulus cells express both connexin 32 and connexin 43 proteins [39, 40]. In porcine and rat cumulus cells, initiation of meiotic resumption is associated with the reduction of the connexin 43 protein level [41, 42]. In the same way, during in vitro maturation of bovine cumulus-oocyte complexes (COCs), the connexin 43-positive gap junctions disappeared whereas connexin 32 became detectable [39].
The aim of the present study was to assess the effects of GH on in vitro cumulus expansion and oocyte maturation in equine and porcine COCs. The cytoplasmic maturation of porcine oocytes was evaluated by in vitro fertilization and development. The expression of GH receptor mRNA was studied in oocytes and cumulus cells of the two species. In order to explain the effect of GH on equine cumulus expansion, the modifications of the expression of proteins implicated in cumulus expansion, i.e., connexins 32 and 43, hyaluronan synthases 1, 2, and 3, were analyzed in equine COCs.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Equine COCs were collected either by transvaginal ultrasound-guided aspiration in the standing mare or from ovaries recovered from mares in a slaughterhouse during the breeding season.
For COC collection by transvaginal ultrasound-guided aspiration, adult cyclic pony mares in good body condition, kept indoors, and fed with concentrates were used in May. Ovarian activity was assessed by routine rectal ultrasound scanning. An injection (i.v.) of 20 mg of crude equine gonadotropin (CEG) was performed when the largest follicle reached 33 mm in diameter to induce ovulation [43]. All follicles larger than 5 mm were punctured at the end of the follicular phase, 24–34 h after induction of ovulation, as previously described [44]. During follicle puncture, mares were sedated with detomidine (Domosedan, 0.6 mg/100 kg body weight (BW) i.v.; Pfizer, Amboise, France) and the rectum was relaxed with propantheline bromide (60 mg/100 kg BW i.v.; Sigma, Saint Quentin Fallavier, France). After follicular fluid aspiration, follicles were flushed with phosphate buffered saline (PBS; Dulbecco, Unipath, Dardilly, France) and heparin (50 IU/ml; LEO S.A., St-Quentin en Yvelines, France) at 37°C. Aspirated fluid from each follicle was examined individually with a stereomicroscope for COC recovery. After puncture sessions, mares received an antibiotic injection (Intramicine, 1 600 000 IU penicillin/100 kg BW and 1.3 g dihydrostreptomycin/100 kg BW i.m.; Rhône Mérieux, Lyon, France).
For COC collection from ovaries recovered from mares in a slaughterhouse, ovaries were obtained immediately after mares were killed and transported to the laboratory within 2 h in 0.9% NaCl at 37°C. The tunica albuginea was removed and all follicles larger than 5 mm were aspirated with an 18
-gauge needle at 30 mm Hg of vacuum pressure. The ovaries were cut into thick sections with a scalpel blade to find other follicles within the ovarian stroma. Follicular fluids were examined with a stereomicroscope for COC recovery.
Equine Cumulus-Oocyte Complexes Maturation and Nuclear Examination
COCs were classified morphologically at recovery as compact or expanded. COCs from follicles <30 mm were washed four times in PBS with gentamycin (50 mg/l, Sigma), once in maturation medium, and cultured individually in 20 µl of maturation medium covered with mineral oil (Sigma) in a humidified atmosphere of 5% CO2 in air at 38.5°C for 30 h. The control maturation medium was TCM 199 with Earle salts (Sigma) supplemented with 5 mg/ml BSA (Sigma), 1 µg/ml estradiol (Sigma), and antibiotics (100 IU/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml fungizone; Gibco, Eragny, France). The control maturation medium was supplemented with either 0.5 µg/ml equine growth hormone (eGH, NIH) or 5 µg/ml equine luteinizing hormone (eLH, NIH).
After culture, cumulus expansion was subjectively assessed as previously described [44]. COCs were rinsed in PBS and stripped of their cumulus cells with a small glass pipette in PBS. Totally denuded oocytes were rinsed in PBS, stained with 1 µg/ml bisbenzimide fluorescent dye (Hoechst 33342; Sigma) in PBS and observed in a drop on a slide with an epifluorescence microscope in order to determine the nuclear stage. Oocytes showing a polar body and two distinct spots of chromosomes were considered to be in metaphase II. They were then rinsed in PBS, frozen dry in liquid nitrogen, and kept at -80°C. The PBS solution containing cumulus cells was centrifuged (13 000 x g, 1 min) in order to obtain a cell pellet. After removal of supernatants, pellets were frozen in liquid nitrogen and stored at -80°C. For further analysis, only oocytes that reached metaphase II after in vitro culture and cumulus cells associated were used.
In order to obtain a mature control for further analysis, expanded COCs collected from preovulatory follicles (
30 mm) were examined at collection as above. Only oocytes that reached metaphase II after in vivo maturation were stored.
In order to obtain an immature control, compact COCs from small follicles (<30 mm) were examined at collection as above. Only oocytes that were at the germinal vesicle stage were stored.
Porcine Cumulus-Oocyte Complex Recovery, Maturation, and Nuclear Examination
The details for porcine COC collection and in vitro maturation have been described previously [45]. Briefly, ovaries were collected from slaughtered prepubertal gilts. COCs with compact cumulus cell mass were washed three times in modified PBS (supplemented with 36 µg/ml pyruvate, 50 µg/ml gentamycine, and 0.5 mg/ml BSA [fraction V, Sigma]), and once in maturation medium. Groups of 50 COCs were transferred into four-well plates, each well containing 500 µl of maturation medium, for 44 h at 39°C in an atmosphere of 5% CO2 in air with maximum humidity. The basal maturation medium was TCM 199 with Earle salts (Sigma) supplemented with 570 µM cysteamine (Sigma). It was supplemented with 0, 0.1, 0.5, or 1 µg/ml porcine growth hormone (pGH, NIH). The control maturation medium was the basal maturation medium supplemented with 10 ng/ml EGF (Sigma) and 400 ng/ml FSH (PRIMUFOL; Rhône Mérieux). The control maturation medium was supplemented with 0 or 0.5 µg/ml pGH.
After culture, cumulus expansion was subjectively assessed as previously described [44]. In each group, 30 COCs were denuded by strong pipetting, rinsed in modified PBS, fixed for 5 min in ethanol:acetic acid:chloroform (6:3:1) and 2 h in ethanol:acetic acid (3:1). They were stained with 10 µg/ml of bisbenzimide fluorescent dye (Hoechst 33342; Sigma) and mounted with glycerol under a cover slip sealed with nail polish. All slides were visualized with an epifluorescence microscope.
For RNA extraction, porcine oocytes were stripped of their cumulus cells with a small glass pipette, stained, observed in a drop, frozen dry, and stored at -80°C as described for equine oocytes. Porcine cumulus cells were centrifuged, frozen dry, and stored at -80°C as described for equine cumulus cells. Only oocytes that reached metaphase II after in vitro culture and cumulus cells associated were used.
Porcine Cumulus-Oocyte Complex Fertilization and Development
The details for porcine COC in vitro fertilization and development have been described previously [45]. After the maturation period, oocytes were denuded by strong pipetting, rinsed three times in modified PBS and once in fertilization medium, and transferred in groups of 25 oocytes in fertilization medium, which was Tris-buffered medium (TBM, [46]). For in vitro fertilization (IVF), oocytes were cultured in 500 µl of TBM with 2 x 105 frozen thawed spermatozoa/ml during 24 h.
After IVF, 30 oocytes per group were fixed in absolute ethanol for 24 h, stained with bisbenzimide, and visualized with an epifluorescence microscope. Normal fertilization referred to zygotes with two polar bodies and two pronuclei. The oocytes containing at least one decondensing spermatozoa were considered as penetrated.
Embryo development took place in 25-µl droplets of NCSU 23aa [47] under paraffin oil in a humidified atmosphere of 5% CO2: 5% O2: 90% N2 at 39°C for 7 days. The cultured embryos were assessed for cleavage 48 h after fertilization under a stereomicroscope. Fetal calf serum (FCS, Sigma) was added (10%) 96 h postinsemination. Blastocysts were counted 7 days postfertilization.
Extraction of Total RNA and Reverse Transcription-Polymerase Chain Reaction
Total RNA was extracted from equine and porcine oocytes and cumulus cells using the TriPure Isolation Reagent Kit (Boehringer Mannheim, Mannheim, Germany) according to the manufacturer's recommendations. Briefly, pools of two cumulus cells pellets or a single oocyte were mixed with 100 µl of reagent and 1 µg of glycogen (Boehringer Mannheim) as a carrier and added to 60 µl of chloroform. After centrifugation, RNA in the upper aqueous phase was recovered, precipitated with isopropanol, washed with ethanol, and dried. After extraction, RNA pellets were resuspended in 4 µl of water and stored at -20°C.
Prior to the reverse transcription (RT) reaction, the RNA samples were incubated for 10 min at 70°C and chilled on ice. The RT reaction was conducted in a final volume of 10 µl, containing 4 µl of extracted RNA, 2 µM oligo(dT)12–18 (Amersham Pharmacia, Orsay, France), 1x reverse transcriptase buffer (Promega, Charbonnière, France), 2.5 mM MgCl2 (Promega), 1.25 mM of each dNTP (Promega), 20 IU of recombinant ribonuclease inhibitor (RNasin, Promega), and 100 IU of Moloney murine leukemia virus reverse transcriptase (M-MLV, Promega). The reaction was performed for 60 min at 37°C, then reverse transcriptase was heat inactivated for 5 min at 95°C.
Polymerase chain reaction (PCR) amplification of growth hormone receptor, beta-actin, and glyceraldehyde-3-phosphate deshydrogenase (GAPDH) cDNA were performed with 4 µl of RT product, in a final volume of 25 µl containing 1x Taq DNA polymerase buffer, 1.5 mM of MgCl2 (Promega), 0.2 µM of each dNTP (Promega), 0.5 µM of each sense and antisense primer (Eurogentec, Nantes, France), and 1 IU of Taq DNA polymerase (Promega). PCR reactions were performed in a thermal cycler (geneamp PCR system 9700; Perkin Elmer, Villebon sur Yvette, France). The initial denaturation was conducted at 94°C for 3 min, followed by 40 cycles of 30 sec at 94°C, 30 sec at primer hybridization temperature (60°C for equine primers or 55°C for porcine primers), and 45 sec at 72°C. Final elongation was performed at 72°C for 3 min. The RT-PCR products were subjected to electrophoresis on a 2% agarose gel containing 0.25 µg/ml ethidium bromide and were visualized by ultraviolet illumination.
Porcine beta-actin primers were based on the mouse sequence of beta-actin [48]. The upstream primer (5'-CTACAATGAGCTGCGTGTGG-3') and the downstream primer (5'-TAGCTCTTCTCCAGGGAGGA-3') predict a 450-base pair (bp) DNA fragment. Primers for equine GAPDH were based on the sequence of the equine GAPDH cDNA published in Genbank (accession number AF157626, [49]). The upstream primer (5'-GTTTGTGATGGGCGTGAACC-3') and the downstream primer (5'-TTGGCAGCACCAGTAGGAGC-3') predict a 280-bp DNA fragment. Primers for pig GH receptor (rGH) were based on the sequence of the porcine rGH cDNA published in Genbank (accession number X54429, [50]). The upstream primer (5'-GCCAATGACGTGTGTGATGG-3') and the downstream primer (5'-TCCCTGCTGGTGTAATGTCG-3') predict a 292-bp DNA fragment. Primers for equine rGH were based on the sequence of the equine rGH cDNA published in Genbank (accession number AF097588, [51]). The upstream primer (5'-GCCTCAACTGGACTCTACTG-3') and the downstream primer (5'-TGCAGTTCATACTCCAGGAC-3') predict a 124-bp DNA fragment.
As negative controls, tubes without RNA or reverse transcriptase were used for RT-PCR. Omitting RNA or reverse transcriptase did not generate any amplified fragments (data not shown).
Gel Electrophoresis and Immunoblotting
One-dimensional (1D) SDS-PAGE was prepared according to the discontinuous buffer system of Laemmli [52] with a 12% separating gel and a 4% stacking gel. Acrylamide-bisacrylamide solution was purchased from Serva GmbH & Co. (Heidelberg, Germany), and other reagents were purchased from Sigma. Each pellet of equine cumulus cells was diluted in 4 µl of electrophoresis buffer (Tris-HCl, 160 mM, pH 6.8, EDTA 10 mM, SDS 10%, beta-mercaptoethanol 10%, glycerol 20%, and bromophenol blue) and boiled for 4 min. Twelve pellets from the same group (immature control, mature control, in vitro maturation in the control medium, with eGH or with eLH) were pooled and loaded on the gel. Electrophoresis was performed at a constant intensity of 25 mA per gel. At the end of migration, gels were electroblotted on hydrophobic polyvinylidene difluoride membranes (PVDF, Amersham Pharmacia Biotech) overnight at 4°C.
Each membrane was used for six successive immunological detections, i.e., hyaluronan synthase (Has) 1, Has 2, Has 3, connexin 43, connexin 32, and actin. The membranes were washed with TBS (10 mM Tris, 150 mM NaCl) containing 0.1% Tween 20 (TBS-Tween 20), incubated for 1 h in blocking solution (5% dry milk, 0.2% Nonidet P40 in TBS), and incubated for 3 h with the primary antibodies. The primary antibodies used were an anti-mouse Has 1 (MC357) rabbit polyclonal antiserum, an anti-mouse Has 3 (MC359) rabbit polyclonal antiserum, an anti-mouse Has 2 (MC285) ligand affinity pure rabbit polyclonal antibody (provided by Dr. McDonald, Mayo Clinic, Arizona), an anti-connexin 43 affinity-isolated rabbit antibody (C-6219 Sigma), an anti-connexin 32 affinity-isolated rabbit antibody (C-3470 Sigma), and a mouse monoclonal anti-actin (sc-8432; Santa Cruz Biotechnology). The membranes were then washed with TBS-Tween 20, incubated for 30 min in blocking solution, and incubated for 1 h with peroxidase-conjugated secondary antibodies. The peroxidase-conjugated secondary antibodies used were a sheep anti-rabbit IgG (Bio-Rad, France) and a goat anti-mouse IgG (Santa Cruz Biotechnology). The ECL+plus western blotting detection system (Amersham Pharmacia Biotech) was used to detect immunoreactive polypeptides with a phosphoimager (STORM, Molecular Dynamics, Leiden, The Netherlands). Signals were quantified using image analysis software (ImagQuant). After each immunological detection, the membranes were stripped of bound antibodies by incubation in stripping buffer (100 mM beta-mercaptoethanol, 2% SDS, 62.5 mM Tris-HCl, pH 6.7) at 50°C for 15 min, and two rinses in TBS (10 mM Tris, 150 mM NaCl) containing 0.1% Tween 20. The amounts of Has 1, Has 2, Has 3, connexin 43, and connexin 32 were analyzed as the ratio to the actin amount. In order to test the specificity of the bands and to adjust the concentration of the antibodies, we previously used membranes loaded with 12 pellets of equine cumulus cells in a lane. Each antibody was tested on a new membrane that was not previously probed with another antibody. Membranes were processed without the primary antibodies to check for the absence of nonspecific bands. The stripped membranes were developed with ECL+plus and exposed to check for the absence of positive signal and to ascertain the efficiency of the stripping procedure.
Statistical Analysis
Due to the low number of equine oocytes collected per day, successive collection days could not be considered as successive sets of trials. Data from all collection days were pooled and considered as one replicate. The Fisher exact test was used to compare maturation rates and cumulus expansion rates of equine oocytes in the three culture media. In the porcine species, four successive trials were performed. The results were analyzed with one-way ANOVA after arcsine transformation of the mean. When allowed, results were then pooled and tested with the Fisher exact test. The nonparametric Kruskal-Wallis test was performed using StatXact 5 software (CYTEL) to compare means of protein expression signals.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
In this experiment, we tested the effect of GH on cumulus expansion during IVM of equine and porcine oocytes. Equine oocytes were collected either by ultrasound-guided aspiration or from slaughtered mare ovaries. Porcine oocytes were recovered from slaughtered gilt ovaries.
All equine COCs collected by transvaginal ultrasound-guided aspiration were compact at collection. After in vitro culture in the control medium, 13% had an expanded cumulus (Table 1). The addition of eGH or eLH in maturation medium significantly increased the percentage of oocytes with cumulus expansion (29% and 58%, respectively, P < 0.05).
|
At the time of collection, 12% of the COCs recovered from slaughtered mares were expanded and 88% were compact. Only compact COCs were used to evaluate the effect of GH and LH in maturation medium on cumulus expansion. After 30 h of culture, the addition of eGH or eLH in maturation medium had no significant effect on cumulus expansion (Table 1).
In porcine oocytes, pGH had no effect on cumulus expansion after 44 h of culture, neither in control nor in basal maturation medium (not shown). In basal medium, cumulus cell disintegration was observed instead of cumulus expansion, and this was not different in the presence or absence of pGH.
Effect of GH in Maturation Medium on Nuclear and Cytoplasmic Maturation
In this experiment, we tested the effect of GH on nuclear maturation of equine and porcine oocytes and on cytoplasmic maturation of porcine oocytes. Cytoplasmic maturation was evaluated by fertilization and early embryo development rates.
Whatever the type of recovery for equine COCs, the addition of eLH in the control maturation medium significantly increased the percentage of oocytes in metaphase II (32% vs. 51% for the total, P < 0.05; Table 2). The addition of 0.5 µg/ml eGH in the control maturation medium increased the percentage of equine oocytes in metaphase II. Due to the low number of oocytes for each type of recovery, the difference was significant when the two types of recovery were gathered (32% vs. 43%, P < 0.05). For porcine oocytes, the addition of 0.5 µg/ml of pGH in the control maturation medium (i.e., in presence of EGF and FSH) had a positive effect on nuclear maturation (89% vs. 78%, P < 0.05; Table 3). The addition of increasing concentrations of pGH in the basal medium had no significant effect compared with the negative control, but the highest dose of pGH (1 µg/ml) increased the percentage of metaphase II porcine oocytes (40% vs. 27%, P < 0.05).
|
|
The positive effects of pGH on nuclear maturation of porcine oocyte when added in control maturation medium was not further observed on cytoplasmic maturation. Indeed, the results of fertilization (Table 4) and of embryo development (Table 5) were not affected by the presence of pGH in the maturation medium. It was interesting to note that between 1% (positive control) and 12% (basal medium plus 0.1 µg/ml GH) of penetrated oocytes were still immature. The percentage of penetrated oocytes (Table 4) and the percentage of cleaved embryos (Table 5) were higher in control than in basal medium. Nevertheless, there was no significant difference in the percentage of blastocysts over cleaved embryos between all groups, indicating that this aspect of cytoplasmic maturation was not affected by the maturation medium in this experiment (Table 5).
|
|
GH Receptor mRNA Expression in Oocytes and Cumulus Cells
The expression of GH receptor mRNA in cumulus cells and in the oocyte was studied in equine and porcine using RT-PCR.
Total RNA was isolated from equine oocytes and cumulus cells after collection, after in vivo maturation, and after in vitro maturation in the control medium supplemented or not with eGH or eLH. Amplification of cDNA with equine GH receptor-specific primers resulted in one PCR product with the expected size of 124 bp (Fig. 1). This GH receptor product was detected in all oocytes and cumulus cells whether they were analyzed after collection, after in vivo maturation, or after in vitro maturation in one of the three media. No PCR products were detected when a water control was used as template for the PCR. As a control for RNA isolation and the production of cDNA, all samples were amplified with equine GAPDH-specific primers. A strong GAPDH product of expected size (280 bp) was detected in all samples (data not shown).
|
Total RNA was isolated from porcine oocytes and cumulus cells after collection and after in vitro maturation in the control medium supplemented or not with pGH. Using RT-PCR with porcine GH receptor-specific primers, GH receptor mRNA was detected in RNA samples isolated from porcine cumulus cells where a single amplified band of the predicted size (292 bp) was obtained (Fig. 2). The GH receptor was amplified in cumulus cells analyzed after collection or after in vitro maturation. As a control, actin product of expected size (450 bp) was detected in all samples. Amplification of cDNA from porcine oocytes resulted in a very faint signal for GH receptor as well as for actin. This could be due to a low quantity of total RNA in single oocytes. However, GH receptor mRNA was detected in some oocytes analyzed after collection (Fig. 2) or after in vitro maturation (data not shown). No PCR products were detected when a water control was used as template for the PCR.
|
Effect of GH in Maturation Medium on Hyaluronan Synthases, Connexin 43, and Connexin 32 Expression
GH stimulated cumulus expansion in equine but not in porcine COCs. Therefore, only equine COCs were used in this experiment. The expressions of hyaluronan synthases (Has) 1, 2, and 3, connexin 43 and 32, and actin were analyzed in equine cumulus cells using gel electrophoresis and immunoblotting (Fig. 3). The antibody raised against Has 1 and the antibody raised against Has 3 revealed three major bands between 44 and 87 kDa. With the Has 1 antibody, the upper and the lower bands were brighter whereas the middle band was faint. With the Has 3 antibody, the middle band was brighter whereas the upper and the lower bands were faint. The connexin 43 antibody revealed two major bands at 43 kDa and 45 kDa and minor bands running between 43 and 45 kDa. The actin antibody revealed one band at 44 kDa. No signal could be detected with the Has 2 antibody and the connexin 32 antibody in our conditions.
|
Twelve cumulus cell pellets from immature controls, from in vitro maturation in the control medium, in the medium supplemented with eGH, in the medium supplemented with eLH, and from mature control were loaded on each lane. The amounts of Has 1, Has 3, and connexin 43 were analyzed as the ratio to the actin amount for each lane. Three gel electrophoreses and immunoblottings were performed, and the mean of the three results was calculated for each protein. The amount of the upper band revealed with the Has 1 antibody was very low in the immature control, whereas it was not different between the three in vitro maturation conditions, or between in vitro and in vivo maturation (Fig. 4). The amounts of the lower band revealed with the Has 1 antibody and the amount of the middle band revealed with the Has 3 antibody were not different between the five conditions. The amount of the upper major band revealed with the connexin 43 antibody was lower in the mature control than in the immature control (P < 0.05; Fig. 4). It was different neither between the three in vitro maturation conditions, nor between in vitro conditions and mature or immature control. The amount of the lower major band revealed with the connexin 43 antibody was not different between the five conditions.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The data presented demonstrate that in vitro maturation in the presence of GH promotes cumulus expansion in equine COCs collected by transvaginal ultrasound-guided aspiration in the standing mare. GH had no effect on cumulus expansion of equine or porcine COCs collected from slaughtered females. The two collection techniques may select two different populations of COCs. In the slaughterhouse, mares are killed at any physiological stage, COC collection from those mares allowed the recovery of a heterogeneous population of COCs, containing expanded and compact cumuluses. On the other hand, COC collection by transvaginal aspiration was performed at the end of the follicular phase, after induction of ovulation, and allowed the recovery of compact COCs only. This technique may select a population of COCs responsive to GH. Moreover, the physiological stage of the mare at the time of COC collection may influence the response of the COCs to the in vitro culture medium, as previously observed [53, 54]. To our knowledge, the influence of GH on cumulus expansion in the equine and porcine species had never been studied until now. In the porcine, EGF and FSH allowed a slight cumulus expansion. In their absence, cumulus disintegration was observed instead of expansion. This reflects the dependence of the process of cumulus expansion on FSH [55]. In the bovine, the influence of GH on cumulus expansion leads to conflicting results. Izadyar et al. [16] showed that 100 and 1000 ng/ml bovine GH induced significant cumulus expansion, whereas Iga et al. [18] observed no cumulus expansion in the presence of 0.1–1 000 ng/ml bovine GH. Both studies were performed with bovine ovaries collected at a slaughterhouse. Cumuli of Blue fox oocytes cultured in a medium supplemented with 10–1 000 ng/ml bovine GH did not expand [22].
In order to elucidate the influence of GH on equine cumulus expansion, we analyzed the process of cumulus expansion during in vivo and in vitro maturation in the presence or in the absence of GH. The process of cumulus expansion involves the deposition of a hyaluronan-rich extracellular matrix by the cumulus cells. The enzyme that is responsible for the synthesis of hyaluronan in COCs has not been characterized in the equine. Recently, three kinds of mammalian genes encoding hyaluronan synthases (Has 1, Has 2, and Has 3) have been identified in the mouse, the human, and the porcine [31–36]. Sequence comparison has indicated low levels (55–71%) of amino acid identity among the three Has within species, in contrast with the high degree (96–99%) of conservation between human and mouse [35], arguing for evolutionary conservation of functionally important residues. We used polyclonal antibodies to mouse Has 1, 2, and 3 [56] to analyze the hyaluronan synthase expressions in the equine cumulus cells. The antibody raised against mouse Has 1 and the antibody raised against mouse Has 3 revealed three major bands between 44 and 87 kDa. No signal could be detected with the Has 2 antibody. The three bands could be due to the expression of three hyaluronan synthases in equine cumulus cells. The expression of hyaluronan synthases in cumulus cells has also been reported in the mouse and the porcine [36, 38]. The molecular masses of these putative equine hyaluronan synthases are consistent with the masses of the mouse Has 1, 2, and 3 that migrate between 46 and 66 kDa [56]. The antibodies raised against mouse Has 1 and Has 3 may cross-react with the three kinds of Has, whereas the antibody against mouse Has 2 may cross-react with none of the equine Hases. In order to confirm these data, further studies are required to assess expression of Has in equine cumulus cells on mRNA level. The amount of the upper band revealed with the Has 1 antibody was very low in cells from immature COCs, and increased during in vitro or in vivo maturation. The correlation of the expression of this Has with the appearance of hyaluronan and cumulus expansion suggest that this protein may represent an active form of equine hyaluronan synthase involved in the synthesis of the hyaluronan-rich extracellular matrix by the cumulus cells during expansion. Similar results were observed in the mouse and the porcine: the mouse Has 2 mRNA is undetectable in COCs before inducing cumulus expansion, and the mRNA levels increase during expansion induced in vivo by hCG injection [38]; the expression levels of porcine Has 2 mRNA in cumulus cells increase during cumulus expansion stimulated in vitro by equine chorionic gonadotropin and porcine follicular fluid [36]. We evaluated the expression of Has in equine cumulus cells after COC in vitro maturation in the control medium supplemented or not with eLH or eGH. The amount of the three bands in equine cumulus cells were not different between the three in vitro maturation conditions. The addition of GH during in vitro maturation does not seem to influence the hyaluronan synthase expressions in equine cumulus cells in our conditions.
The cumulus expansion is also accompanied by modifications of gap junctions within cumulus cells. Gap junctions are channels formed by hexametric structures consisting of connexin molecules. To our knowledge, the expression of connexin molecules in granulosa or cumulus cells has never been studied in the equine. Immunoblotting analysis of extracts of equine cumulus cells revealed that the anti-connexin 43 antibody recognized major and minor bands between 43 and 45 kDa. These bands most likely represent various states of phosphorylation of connexin 43. In fact, the existence of phosphorylated forms of connexin 43 in ovarian follicles has been previously reported in several species. In the rat, several studies demonstrate that the connexin 43 gap junction protein is present in multiphosphorylated forms: the 43-kDa band represents the nonphosphorylated form, while the bands with the slightly retarded electrophoretic mobility represent the phosphorylated forms [57–59]. In porcine, immunoblot analysis revealed the presence of two immunoreactive bands of 43 and 45 kDa [60, 61] or three bands of 43, 45, and 47 kDa [42]. The 43-kDa band is a nonphosphorylated band, whereas the 45- and 47-kDa bands are phosphorylated forms of this protein. In our study, the major band at 43 kDa probably represents a nonphosphorylated form of equine connexin 43, the minor bands running between 43 and 45 kDa and the major band at 45 kDa probably represent phosphorylated forms. Further studies with treatment of the samples with phosphatases would confirm these results. Our data show that equine cumulus cells express connexin 43 proteins, as previously observed in bovine, porcine, and mouse cumulus cells [39, 40, 42]. Despite the visual effect on the Western blot, the ratio proves that the amount of the 45-kDa band decreased during COC in vivo maturation, whereas the amount of the 43-kDa band did not vary. These data demonstrate a positive correlation between meiosis resumption and reduction of connexin 43 in gap junctional communication in cumulus cells. They suggest that the balance of the phosphorylated and the nonphosphorylated forms changes during equine COC maturation. The question about the role of phosphorylation of connexin 43 remains to be answered. These results are consistent with data from other species. Immunoblotting analysis of porcine cumulus cells revealed that, during in vitro maturation, the intensity of 43 and 45 kDa connexin 43 bands was reduced [42]. Immunofluorescence analysis of bovine COCs showed that the connexin 43-positive gap junctions disappeared during in vitro culture [39]. In the present study, the expression of connexin 43 was evaluated in equine cumulus cells after COC in vitro maturation in the control medium supplemented or not with eLH or eGH. The amounts of the major bands at 43 and 45 kDa were not different between the three in vitro maturation conditions. The addition of GH during in vitro culture does not seem to influence the connexin 43 expression or phosphorylation in equine cumulus cells.
In equine cumulus cells, no signal could be detected with the connexin 32 antibody in our conditions. The expression of connexin 32 mRNA was undetectable in porcine cumulus cells [62], while transcripts of connexin 32 were detected in mouse cumulus cells [40] and connexin 32 protein expression was demonstrated in mouse and bovine cumulus cells [39, 40]. The lack of signal in equine cumulus cells with the connexin 32 antibody may be related to too low amounts of connexin 32 that could not be detected in our conditions. In fact, in another immunoblot analysis using very large amounts of equine cumulus cells, the connexin 32 antibody revealed one faint band at 32 kDa.
Our data show that the addition of eLH or eGH in maturation medium significantly increased the percentage of equine oocytes in metaphase II. The majority of protocols for maturation of equine oocytes include the addition of LH. Moreover, equine gonadotropins are superior to gonadotropins from other species in inducing equine oocyte maturation [63]. It seems reasonable that the oocyte in vitro would require exposure to LH, because this is the hormone present at the time of in vivo maturation. The influence of GH on equine oocyte maturation had never been studied until now. The positive effect of eGH on nuclear maturation of equine oocytes is in agreement with the results observed in other species. Addition of 100 ng/ml ovine GH to the culture medium increases the percentage of germinal vesicle breakdown in cumulus-enclosed rat oocytes [21]. Addition of 10–1000 ng/ml bovine GH increases the percentage of metaphase II in bovine COCs [12, 18], and leads to a decrease of the percentage of germinal vesicle and an increase of the percentage of metaphase II in Blue fox oocytes [22]. In the bovine, GH not only promotes nuclear maturation but also improves cytoplasmic maturation of the oocytes, the latter reflected by the enhanced migration of cortical granules, sperm aster formation, and blastocyst formation [16–18]. In the equine, further studies are required to evaluate the influence of GH on cytoplasmic maturation. An improvement of in vitro embryo production in the equine would be of great importance because major problems are encountered in this species.
In the pig, a number of attempts have been conducted to improve oocyte maturation in vitro, including the addition of growth factors to the culture media. EGF increases both nuclear [64, 65] and cytoplasmic maturation [45, 66, 67]. To our knowledge, the effects of GH on porcine oocyte maturation were studied only once [20], and the results suggested an acceleration of oocyte maturation. We showed here that GH improved the nuclear maturation rate of porcine oocytes in vitro. It was more effective in the presence of EGF and FSH than alone. This result may reflect a synergistic action between GH and gonadotrophs, as reported in human [68, 69] and in pig [4] granulosa cell steroidogenesis. The synergy between GH and gonadotrophs may also reflect an upregulation of gonadotroph receptors by GH or the upregulation of GH receptors by gonadotroph-induced cAMP [1, 70]. Indeed, at least in sheep, adequate ovarian GH receptor gene expression requires pituitary gonadotrophs [71].
The positive effects of GH on nuclear maturation of porcine oocytes was not associated with an improvement of cytoplasmic maturation because the fertilizability of the oocytes was not affected nor was the developmental competence of the zygotes obtained after IVF.
The precise mechanism by which GH exerts its effects on the COCs is still controversial. In vitro maturation of rat COCs in the presence of GH and anti-IGF-I antibodies showed that, in this species, the effect of GH is mediated by IGF-I [21]. When bovine COCs were cultured in the presence of GH and anti-IGF-I antibodies, conflicting results were obtained. Iga et al. showed that the stimulatory effects of GH are mediated via IGF-I [29], whereas Izadyar et al. concluded that GH-induced oocyte maturation is not mediated by IGF-I [12], but is mediated by the cAMP signal transduction pathway [30]. In order to elucidate the mechanism of GH-induced nuclear maturation in equine and porcine oocytes, we first investigated whether equine and porcine COCs contain GH receptor. The expression of GH receptor mRNA was detected in cumulus cells and in the oocyte in both species. Although the presence of mRNA is indicative but not a proof for a functional receptor, together with the observed responses of GH on oocyte maturation, it can be assumed that equine and porcine COCs do possess a functional GH receptor. These results are consistent with observations in other species. In sheep, mRNA for GH receptor were detected in oocytes [13]. Expression of GH receptor in rat cumulus cells and oocytes has been reported [10]. Both immunoreactive protein and mRNA were found in bovine cumulus cells and oocytes [11, 12, 28]. The presence of GH receptor mRNA in equine and porcine denuded oocytes suggest the possibility for the GH to act directly on the oocyte. However, the lack of effect observed when GH was added to cultures of rat and bovine denuded oocytes indicates that the action of GH is mediated by cumulus cells in these species, in spite of the presence of GH receptor in oocytes. Further studies are required to investigate the importance of cumulus cells for GH-promoted oocyte maturation in equine and porcine species. The first step would be to confirm that translation of the GH receptor mRNA takes place in the oocyte, and that GH receptor protein is expressed. Using radiolabeled GH, Quesnel [14] detected binding sites for GH in porcine oocytes. The method, however, did not allow unequivocal distinction between binding to GH receptor and binding to GH-binding proteins (GHBPs). The second step would be to investigate the effect of GH when added to cultures of equine and porcine denuded oocytes.
In conclusion, our results confirm the importance of GH in the ovarian physiology. A better understanding of the mechanism by which GH stimulates the oocyte can have important implications in the use of GH for in vitro embryo production in domestic mammals.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
2 Correspondence: Ghylène Goudet, Unité de Physiologie de la Reproduction et des Comportements, I.N.R.A., 37380 Nouzilly, France. FAX: 33 2 47 42 77 43; goudet{at}tours.inra.fr
Received: 17 January 2003.
First decision: 10 February 2003.
Accepted: 8 May 2003.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
