The involvement of growth hormone in equine oocyte maturation, receptor localization and steroid production by cumulus–oocyte complexes in vitro

Research in Veterinary Science 95 (2013) 667–674

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The involvement of growth hormone in equine oocyte maturation, receptor localization and steroid production by cumulus–oocyte complexes in vitro G.R. Pereira a,e,⇑, P.L. Lorenzo b, G.F. Carneiro a, B.A. Ball a, S. Bilodeau-Goeseels c, J. Kastelic c, L.M.C. Pegoraro d, C.A. Pimentel e, A. Esteller-Vico b, J.C. Illera b, G.S. Granado b, P. Casey f, I.K.M. Liu a a

Department of Population Health and Reproduction, School of Veterinary Medicine, University of California, Davis, CA 95616, USA Animal Physiology Department, Veterinary School, Universidad Complutense de Madrid, Madrid 28040, Spain c Agriculture and Agri-Food Canada, Lethbridge Research Center, Lethbridge, AB T1J 4B1, Canada d Animal Reproduction Laboratory, Temperate Climate Research Corporation, EMBRAPA, Pelotas, RS 96001-970, Brazil e Department of Animal Pathology, Animal Reproduction Laboratory, School of Veterinary Medicine, Federal University of Pelotas, Capão do Leão s/n, Mailbox 354, Pelotas, RS 96010-900, Brazil f Research Centre in Reproductive Medicine, Faculty of Medicine and Health Science, The University of Auckland, Auckland, New Zealand b

a r t i c l e

i n f o

Article history: Received 13 September 2012 Accepted 30 June 2013

Keywords: Oocyte maturation Growth hormone Steroidogenesis Growth hormone receptor Horse

a b s t r a c t The objectives of this study were to evaluate the effects of equine growth hormone (eGH) on nuclear and cytoplasmic maturation of equine oocytes in vitro, steroid production by cumulus cells, and expression and subcellular localization of eGH-receptors (eGH-R) on equine ovarian follicles. Cumulus–oocyte complexes (COCs) were recovered by aspirating follicles <30 mm in diameter from abattoir-derived ovaries. The COCs were morphologically evaluated and randomly allocated to be cultured in either a control maturation medium or supplemented with 400 ng/mL eGH, for 30 h at 38.5 °C in air with 5% CO2. The COCs were stained with 10 lg/mL propidium iodide and 10 lg/mL fluorescein isothiocyanate-labeled Lens culinaris agglutinin. Chromatin configuration and distribution of cortical granules were assessed via confocal microscopy. Compared to control, COCs incubated with eGH had: more oocytes that reached metaphase II (35/72, 48.6% vs. 60/89, 67.4%, respectively; P = 0.02); greater concentrations of testosterone (0.21 ± 0.04 vs. 0.06 ± 0.01 ng/mL; P = 0.01), progesterone (0.05 ± 0.01 vs. 0.02 ± 0.00 ng/mL; P = 0.04), and oestradiol (76.80 ± 14.26 vs. 39.58 ± 8.87 pg/mL; P = 0.05) in the culture medium, but no significant differences in concentration of androstenedione. Based on Real Time RT-PCR analyses, expression of the eGH-R gene was greater in cumulus cells and COCs at the start than at the end of in vitro maturation. Positive immunostaining for eGH-R was present in cumulus cells, the oocytes and granulosa cells. In conclusion, addition of eGH to maturation medium increased rates of cytoplasmic maturation and had an important role in equine oocyte maturation, perhaps mediated by the presence of eGH-R in ovarian follicles. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction Routine in vitro fertilization (IVF) procedures in the horse have been hampered by factors, including difficulties related to oocyte maturation and maintenance of the developing embryo (Hinrichs, 2010). The proportion of equine oocytes maturing in vitro has

⇑ Corresponding author at: Department of Animal Pathology, Animal Reproduction Laboratory, School of Veterinary Medicine, Federal University of Pelotas, Capão do Leão s/n, Mailbox 354, Pelotas, RS 96010-900, Brazil. Tel.: +55 55 99972240; fax: +55 53 32731937. E-mail addresses: [email protected], [email protected], [email protected] (G.R. Pereira). 0034-5288/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.rvsc.2013.06.024

varied widely among laboratories, despite important advances following reports of successful intracytoplasmic sperm injection (Lazzari et al., 2002; Galli et al., 2007) and cloning in the horse (Galli et al., 2003; Lagutina et al., 2005). Successful IVF is not yet well established in the horse. Efficient techniques for in vitro oocyte maturation are crucial to define culture conditions that promote equine oocyte maturation competence and to provide new strategies to improve the efficiency of assisted reproductive technologies in the horse (Choi et al., 2013; Hinrichs, 2012). Growth hormone (GH) is an important regulator of reproduction in female mammals and has key important roles in ovarian function, including ovarian follicular growth and steroidogenesis (Yoshimura et al., 1993; Bachelot et al., 2002). It is well established

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that GH, a polypeptide hormone, is considered the primary regulator of somatic growth and is produced mainly in the anterior pituitary by somatotropes (Glasscock et al., 1991; Borski et al., 1996). It acts directly through its own receptor, as well as indirectly by inducing production of insulin-like growth factor-I (IGF-I), an important effector peptide that works primarily as a circulatory second messenger of GH in the somatomedin axis (Berelowitz et al., 1981; Rotwein et al., 1994). Furthermore, GH acts independently of IGF-I in mouse follicular development, as the number of follicles per ovary was markedly reduced in mice lacking GH receptor (GH-R) and GH binding protein, despite adequate exogenous gonadotrophins (Apa et al., 1994; Bachelot et al., 2002). Thus, it is well established that GH, the GH-R and IGF-I, all have critical roles in mediating cell growth (Liu and LeRoith, 1999; Lupu et al., 2001). Exogenous equine growth hormone (eGH) enhanced equine oocyte nuclear maturation and cumulus oocyte complex (COC) expansion, manifested by increases in the percentage of equine oocytes reaching metaphase-II (M-II) relative to the control (44 vs. 32%, respectively; Marchal et al., 2003). Furthermore, in that experiment, cytoplasmic maturation (assessed with IVF of porcine oocytes) was not affected by the addition of GH in vitro. However, adding recombinant bovine GH (r-bGH) to gonadotrophin-supplemented in vitro maturation (IVM) medium significantly improved early embryo development and increased blastocyst production in cattle (Izadyar et al., 1996, 1998, 2000). In the rhesus macaque, the addition of recombinant human GH (r-hGH) during IVM did not affect nuclear maturation, although r-hGH improved subsequent embryo development in this species (de Prada and VandeVoort, 2008). Studies in our laboratory demonstrated that more equine oocytes resumed meiosis in the presence of eGH and IGF-I in the IVM system after 30 h of culture in serum-free culture medium (Pereira et al., 2006, 2012). Various cytoplasmic changes were reported during maturation of equine oocytes (Grondahl et al., 1995; Goudet et al., 1997; Carneiro et al., 2002). Although migration of cortical granules (CGs) has been assessed as an indicator of cytoplasmic maturation, the ultimate sign of cytoplasmic maturation is the ability of the oocyte to undergo fertilization and develop into a viable embryo (Ducibella et al., 1988; Ferreira et al., 2009). In the absence of an efficacious conventional IVF technique for the horse, a validation study (e.g. using ICSI) is needed to determine the value of migration of the CGs as a marker of cytoplasmic maturation. Carneiro et al. (2002), after reviewing the literature, decided to use migration of CGs as one potential signal for equine cytoplasmic maturation. In mice and pigs, proliferation of granulosa cells reflected the influence of the oocyte participating in steroidogenesis and promoting maintenance of progesterone (P4) production (Morbeck et al., 1993; Vanderhyden and Tonary, 1995). Lorenzo et al., (1997a) reported the influence of IGF-I on steroid production by rabbit COCs during IVM and confirmed the influence of steroiddependent growth factors on COCs. Estradiol (E2) increased steroidogenesis by equine granulosa cells, perhaps by interacting with IGF-I, when insulin was added to the medium used to culture small and medium follicles (Davidson et al., 2002). Mares induced to ovulate with a crude equine gonadotrophin preparation had decreased P4 concentrations, but increased estradiol-17b concentrations in follicular fluid, emphasizing the role of steroidogenesis on oocyte competence at the time of ovulation (Caillaud and Gerard, 2009). Based on in vitro studies in rabbits and pigs, it has been suggested that GH increased intra-ovarian production of IGF-I by stimulating follicular development, oestrogen production, and oocyte maturation (Hsu and Hammond, 1987; Yoshimura et al., 1996). There are few reports on the effects of exogenous eGH on the IVM of equine oocytes and on steroidogenesis by their COCs (Pereira et al., 2006).

The expression of GH-R in rat cumulus cells and oocytes promotes GH binding of the ligand to the receptor to stimulate oocyte maturation (Lobie et al., 1990). Furthermore, based on the presence of GH-R mRNA in equine denuded oocytes, eGH may act directly on the oocyte (Marchal et al., 2003). Thus, eGH-R may mediate a positive effect when eGH is used in culture media during equine oocyte maturation. The identification of eGH-R by immunohistochemistry (IHC) enables identification of a protein constituent in situ, thereby providing important information regarding eGH-R mechanisms associated with oocyte maturation in the horse. The influence of eGH on cytoplasmic maturation and on steroidogenesis in equine COCs has been investigated by our group (Pereira et al., 2006, 2012). However, further information on the eGH-R localization and expression may provide a foundation for strategies to enhance oocyte development in the horse. The objectives of this study were to evaluate the effects of eGH on equine nuclear and cytoplasmic maturation and on steroidogenesis to improve competence of equine oocytes matured in vitro. Additionally, we evaluated the presence of eGH-R on equine ovarian structures including follicular wall, oocytes and COCs by IHC, and expression of eGH-R in COCs and cumulus cells by real time RT-PCR, before and after IVM.

2. Materials and methods All chemicals used were purchased from Sigma Chemicals Company (St. Louis, MO, USA), unless otherwise indicated.

2.1. Collection and culture of cumulus–oocyte complexes Equine ovaries were obtained during the physiological breeding season from abattoirs within 20 min of the two research centers (Lethbridge Research Centre/Canada and EMBRAPA Research Center/Brazil). At each location, ovaries were collected after slaughter and the protocols for oocyte collection, culture and transport were identical. Ovaries were immediately transported in 0.9% NaCl with 100 IU/mL penicillin and 50 lg/mL streptomycin sulfate to the laboratory in an insulated container at 30 °C. The interval between ovary retrieval and culture of COCs ranged from 2 to 3 h. Each follicle <30 mm in diameter was aspirated with an 18-G needle connected to a 35 mL syringe. During aspiration, a scraping motion within the follicle was performed with the needle, to enhance oocyte recovery. The follicular fluid from each follicle aspirated was placed into 100-mm Petri dishes and observed under a stereomicroscope for the presence of COCs. Morphology and structural integrity of each COC was evaluated and classified as compact, expanded or degenerated oocytes, as described by Hinrichs et al. (1993). The basal medium was TCM-199 supplemented with 0.1% bovine serum albumin (BSA), 100 IU/mL penicillin, and 50 lg/mL streptomycin sulfate. The medium was filtered through a 0.22 lm pore diameter filter into 50 mL centrifuge tubes and allowed to equilibrate for at least 1 h at 38.5 °C under 5% CO2 in air prior to use. Recombinant eGH was manufactured by BresaGen Limited (Adelaide, South Australia, Australia). Following morphological classification, selected COCs were washed four times in the basal medium and randomly allocated to either a control group (no additives) or supplementation with 400 ng/mL of eGH containing 500 lL of maturation medium according to Pereira et al. (2012). Numbers of COCs submitted to culture ranged from 8 to 12 for each treatment group. After 30 h of maturation at 38.5 °C in air with 5% CO2, cumulus cells were removed with a 0.1% hyaluronidase solution in PBS.

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2.2. Oocyte fixation and staining The zona pellucida was removed from the cumulus cells using 0.1% pronase in PBS. Zona-free oocytes were fixed in 2% paraformaldehyde for at least 1 h at 5 °C, and subsequently stored in blocking solution containing PBS, 0.2% sodium azide, 1 mg/mL polyvinyl alcohol, and 100 mM glycine (Simerly and Schatten, 1993). Zona-free oocytes were stored in blocking solution and transported in an insulated container at 5 °C to the University of California – Davis for staining and confocal analyses. Oocyte staining and laser-scanning confocal microscopy analysis were performed as previously described (Carneiro et al., 2002). Staining with 10 lg/mL propidium iodide was used to evaluate nuclear status of the oocytes. Cortical granules were stained with 10 lg/mL fluorescein isothiocyanate-labeled lens culinaris agglutinin (Vector) and migration of cortical granules from the centre to the periphery of the oocyte was used as an indicator of cytoplasmic maturity, as described (Damiani et al., 1996; Carneiro et al., 2002). Zona-free oocytes were placed on slides with a space between the coverslip and the slide filled with Vectashield antifade mounting medium (Vector). 2.3. Assessment of oocyte maturation Nuclear and cytoplasmic maturation evaluations were performed by laser-scanning confocal microscopy using a Bio-Rad MRC 1024 ES microscope equipped with a Krypto-argon ion laser. Migration of cortical granules and chromosome visualization were identified simultaneously and recorded with laser scanning confocal microscopy. The presence of chromosomes in the metaphase plate configuration was an indicator of complete nuclear maturation, whereas migration of the CGs to the oocyte periphery was used as an indicator of complete cytoplasmic maturation (Carneiro et al., 2002; Pereira et al., 2012). Zona-free oocytes were classified as immature when clusters of CGs were distributed homogeneously throughout the cytoplasm and no chromosomes aligned at the metaphase plate. Zona-free oocytes at metaphase I (M-I) were characterized by the presence of chromosomes peripherally in the ooplasm and CGs randomly distributed in the cortex cytoplasm, whereas those with chromosomes aligned at the metaphase plate and/or evidence of cortical granule migration at the periphery of the oocyte were both classified as metaphase II (M-II). Atypical cytoplasmatic structures associated with lost chromosomes and accumulations of an amorphous mass near the condensed chromatin were regarded as degenerated oocytes. Each zona-free oocyte was observed with optical sections at 0.1–2.0 lm intervals. 2.4. Immunohistochemistry analysis Immunohistochemistry was used to determine the presence of the eGH-R protein in equine follicles, oocytes, and pituitary tissues. Paraffin sections (5 lm thick) containing an equine oocyte attached to the follicular wall (n = 5) and equine follicles (n = 5) derived from follicles 10–30 mm in diameter were mounted onto poly-L-lysine coated slides and treated by the streptoavidin–biotin method (Histostain-Plus Kit: Zymed). Furthermore, equine oviductal (n = 2) and pituitary (n = 2) tissues were also used for detection of eGH-R. A 10% non-immune goat serum blocking solution in PBS was used as a conjugate for primary antibody dilutions. Conjugate only was used as a negative control, whereas pituitary tissue samples were used as a positive control. Negative controls, using preimmunized rabbit serum, were processed for sampled sections. Primary monoclonal mouse-anti GH-R antibodies; Mab 263 (MCA1555; AbD Serotec) at 1:20 dilution were incubated overnight at 4 °C in a humidified chamber. It is noteworthy that Mab 263 reacts against an epitope located within the first disulphide

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loop of the rat and rabbit GH-R, which is highly conserved among species (Gobius et al., 1992). Samples were later incubated with biotinylated goat anti-rabbit IgG secondary antibody (1:1200 dilution) for 30 min, followed by 30 min incubation with the streptoavidin–biotin–peroxidase complex at 1:400 dilution. Tissue sections were then incubated for 10 min with AEC Single Solution Chromogen (85-6543: Zymed) and counterstained with Hematoxylin (H-3403: Vector) for 3 s. Slides were washed in ddH2O for 5 min before mounting with Faramount Aqueous Medium (S-3025; Dako) until dry. Sampled slides were evaluated using a Leica DMRB microscope and images were saved using QCapture imaging software (QImaging). 2.5. Assessment of steroid production Following removal of the COCs, the maturation medium was collected and stored at 20 °C to measure steroid production by enzyme immunoassay (EIA; Elisa Aid, Eurogenetics) to investigate the influence of eGH on production of progesterone (P4), estradiol17b (E2), androstenedione (A4) and testosterone (T4), as described (Lorenzo et al., 1997b). General EIA procedures, such as immunization, determining the titre of antiserum, and purification of antibodies for steroid production were conducted using polyclonal antibodies produced in adult New Zealand White rabbits. Testosterone 3-carbomethyloxime:BSA, androstenedione 3-carboxymethyloxime:BSA, 6-keto-17estradiol 6-carboximethyloxime:BSA, and 11-hydroxiprogesterone 11 hemisuccinate:BSA were used as immunogens for T4, A4, E2, and P4, respectively (Illera et al., 1997; Lorenzo et al., 1997b). Hormone determinations were done, in duplicate, in culture media without previous extraction. The lower detection limits were 2.0 pg/well (100 lL) for E2, 13.2 pg/well for P4, 2.3 pg/well for A4, and 11.4 pg/well for T4. Intra- and inter-assay coefficients of variation percentages (% CV) were <7.8% in all cases. The standard dose response curve covered a range between 0 and 1 ng/well. 2.6. Real-time reverse transcription polymerase chain reaction (qRT-PCR) Prior to IVM and at the end of the 30 h culture with eGH, cumulus cells were removed with 0.1% hyaluronidase solution. Pools of 15 compact COCs selected prior to IVM and denuded oocytes after IVM per se were subjected to qRT-PCR analysis, which was used to quantify the relative transcript levels of GH-R (Leutenegger et al., 1999). Briefly, primers and a TaqMan probe for the equine GH-R TaqMan PCR system were designed using Primer Express software (ABI; Applied Biosystems). The GH-R TaqMan probe (50 -FAMCAGATATCCAAGTGAGGTGGGAACCACCA-TAMRA-30 ) spanned the junction of two consecutives exons covered by the primers (forward primer 50 -AACATCAGTTTAACCGGGATTCAT-30 , and reverse primer 50 -GATCACAGCTCCGGAAGCAG-30 ) to restrict the TaqMan PCR specificity to cDNA without detecting genomic DNA contamination. Primers for eGH-R precursor were based on a cDNA sequence published in GenBank (Accession No.: AF097588). The TaqMan PCR systems were validated using 2-fold dilutions of cDNA testing positive for target genes. Dilutions were analyzed in triplicate and a standard curve plotted against dilutions. In brief, the housekeeping gene, glyceraldehyde-3-phosphate-dehydrogenase (GAPDH) was used to normalize the comparative (CT) values of the target genes (DCT). The DCT was calibrated against the weakest signal within each target gene. The relative linear amount of target molecules, relative to the calibrator, was calculated by 2DDCT. Therefore, all gene transcription was expressed as an n-fold difference relative to the calibrator. Final quantification was done using the CT method and reported as relative transcription or the n-fold difference relative to a calibrator cDNA (i.e., lowest target gene

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transcription) after normalization of the GH signal to the endogenous control (GAPDH). 2.7. Statistical analysis Percentages of oocyte maturation after eGH treatment groups were analyzed by Chi-square (v2) test, using PROC CATMOD

(SAS; SAS Institute). Steroid hormone concentrations in culture media were compared between treatments with one-way analysis of variance (ANOVA; JMP, SAS Institute). Data were normally distributed as assessed using the Shapiro–Wilk test and were log-transformed before analysis (to normalize distributions). Data are expressed as means ± SEM. The level of significance was set at P < 0.05.

Fig. 1. Equine oocytes evaluated with confocal microscopy after 30 h of incubation in medium supplemented or not with 400 ng/mL equine growth hormone. Cortical granules (green) localization and distribution on serial optical sections in equine oocytes. Images are representative of confocal investigations performed on all examined oocytes. (A1–12) Immature oocyte with cortical granules distributed in the cortex of the cytoplasm and no visible chromatin. (B1–12) Oocyte at metaphase-I (M-I) with condensed chromosomes (red: B3–9) in the ooplasm and the CGs randomly distributed in the cortex cytoplasm. (C1–12) Oocyte at metaphase-II (M-II) note chromosomes (red) aligned along the metaphase plate (C8–11) and CGs released to the periphery after oocyte maturation. (D) Immature oocyte with CGs distributed in the cortex of the cytoplasm and no visible chromatin. (E) Oocyte at M-II with chromosomes aligned along the metaphase plate and CGs released to the periphery. Scale bar A–C represents 70 lm and D and E represents 30 lm. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

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3. Results Immature, M-I and M-II oocytes are shown (Fig. 1) and were classified as previously described (Carneiro et al., 2002). Following incubation with 400 ng/mL eGH, 67.4% oocytes resumed meiosis (M-I + M-II) compared to 48.6% in the control group (P = 0.02; Table 1). The proportion of equine oocytes at germinal vesicle (GV) stage was higher in the control group (38.9%) compared to the eGH group (24.7%; P = 0.02). The culture medium of COCs incubated with eGH had higher concentrations of T4, P4 and E2 compared to the culture medium of control oocytes (Table 2; P < 0.05).

Immunostaining for eGH-R was evident in cumulus and granulosa cells, as well as the oocyte; these signals were most prominent in the oocyte and cumulus cells of developing follicles (Fig. 2A and B). Positive immunostaining was also detected in theca interna cells in the follicular wall (Fig. 2C). Immunoreactive eGH-R in the cytoplasm of the anterior pituitary somatotropes cells was strongly stained after incubation with monoclonal antibody 263 (Fig. 2D). Negative control tissues were incubated with IgG at the same concentration as the primary antibody (Fig. 2A–C inset). Positive control samples of equine pituitary had no positive immunostaining in the absence of primary antibody (Fig. 2D inset).

Table 1 The effects of equine growth hormone at 400 ng/mL on nuclear and cytoplasmic maturation of equine oocytes in vitro after 30 h of culture. Culture groups

Control eGH

No. oocytes cultured

114 163

No. oocytes evaluated

72 89

No. (%) at each maturation stage Degenerated

Immature

M-I

M-II

M-I + M-II

9 (12.5)a 7 (7.9)a

28 (38.9)a 22 (24.7)b

13 (18.0)a 23 (25.8)a

22 (30.6)a 37 (41.6)a

35 (48.6)a 60 (67.4)b

Equine growth hormone (eGH), metaphase-I (M-I), metaphase-II (M-II). a,bWithin a column, percentages without a common superscript differed (P = 0.02). Data were pooled from 11 replicates.

Table 2 Mean ± SEM concentrations of steroid hormones in the culture media of equine cumulus oocyte complexes after 30 h of culture under control conditions or in the presence of 400 ng/mL equine growth hormone. Culture groups

No. COCs cultured

T4/COCs (ng/mL)

P4/COCs (ng/mL)

E2/COCs (pg/mL)

A4/COCs (ng/mL)

Control eGH

114 163

0.06 ± 0.01a 0.21 ± 0.04b

0.02 ± 0.00c 0.05 ± 0.01d

39.58 ± 8.87e 76.80 ± 14.26f

0.08 ± 0.02g 0.08 ± 0.01g

Equine growth hormone (eGH), testosterone (T4), progesterone (P4), estradiol (E2), androstenedione (A4), and cumulus oocyte complexes (COCs). a–fWithin a column, means without a common superscript differ: a vs. b; P = 0.01, c vs. d; P = 0.04, and e vs. f; P = 0.05. Data were pooled from 11 replicates.

Fig. 2. Localization of eGH-R by immunohistochemistry in ovarian follicular structures. As a negative control, sections were incubated with IgG at the same concentration as the primary Mab 263 antibody (inset). Equine anterior pituitary tissue was used as a positive control for eGH-R detection. (A) Positive immunostaining for eGH-R in the equine oocyte (Oo). (B) Immunoreactivity for eGH-R in cumulus oocyte complexes (COCs). (C) Immunoreactivity for eGH-R in the theca interna. (D) Immunoreactivity for eGH-R in somatotropes. Oo, Oocyte; ZP, Zona pellucida; GC, Granulosa cell; T, Theca interna; eGH-R; Equine growth hormone receptor. ⁄ Presence of eGH-R (red stain). Bar represent 20 lm.

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Fig. 3. Relative gene expression of eGH-R in cumulus cells from 15 COCs and cumulus oocyte complexes (pool of 15 COCs) before and after culture for 30 h with eGH in vitro. Gene expression was measured by qRT–PCR using TaqMan and analyzed with the delta Ct method. Relative gene transcription was reported as an n-fold difference relative to a calibrator cDNA.

Culture of COCs with eGH for 30 h resulted in an 11-fold down-regulation of the eGH-R mRNA compared to COCs collected prior to IVM. Based on quantification by Taq-Man PCR, relative gene transcription for eGH-R was higher in cumulus cells and COCs at the start of in vitro culture than after 30 h of incubation (Fig. 3).

4. Discussion and conclusions In the present study, 67.4% of equine oocytes that matured in vitro in the presence of eGH resumed meiosis after IVM, whereas Marchal et al. (2003) reported a 43% increase in nuclear maturation of M-II oocytes. The migration of CGs has been assessed as an indicator of cytoplasmic maturation of equine (Carneiro et al., 2002), bovine (Damiani et al., 1996) and murine (Ducibella et al., 1994) oocytes, where density significantly decreased in the central area compared with the periphery as maturation proceeded, although the ultimate sign of cytoplasmic maturation was the ability of the oocyte to undergo fertilization and develop into a viable embryo (Ducibella et al., 1994; Ferreira et al., 2009). Recently, we reported a higher maturation rate (73.3%) when eGH was added to the maturation medium with IGF-I; therefore, the cytoplasmic pattern during oocyte maturation became more evident when we used eGH in combination with IGF-I, indicating its role in promoting development of equine oocytes cultured in vitro (Pereira et al., 2012). This study was also designed to determine the effect of eGH on steroidogenesis under serum-free culture conditions, thereby eliminating the influence of unknown serum factor(s). In cattle, although T4 is secreted in low concentrations by COCs during in vitro culture, its potential involvement in cumulus expansion makes it a prerequisite for bovine oocyte maturation (Wang et al., 2006). In the present study, eGH-treated COCs produced significantly more T4 than the control group; therefore, we inferred that T4 might have an important role in equine oocyte maturation. In vivo, intrafollicular T4 concentrations did not vary with follicle diameter in the mare, although growth of the dominant follicle was accompanied by increased intrafollicular concentrations of estradiol-17b and P4 (Belin et al., 2000). Furthermore, exogenous T4 increased E2 production, indicative of the presence of an active aromatase enzyme system in equine granulosa cells (Sirois et al., 1991). In the present study, E2 concentrations were slightly increased after incubation with eGH, in contrast with studies in Aromatase Anockout Mouse, where E2 secretion appeared not to be essential for maintenance of healthy immature oocytes in vivo (Hu et al., 2002; Huynh et al., 2004).

The increase of P4 synthesis in the present study was consistent with progesterone-enhanced bovine oocyte IVM and development (Karlach, 1987). In mares, P4 production was higher in culture of granulosa cells from mares in late oestrus compared to those in early oestrus (Sirois et al., 1990). Furthermore, increased E2 secretion in the present study was supported by Mingoti et al. (2002) who reported that bovine COCs secreted high concentrations of E2 and P4 during in vitro maturation and cumulus expansion. Researchers reported higher expression of steroidogenic acute regulatory (StAR) mRNA in IVM of macaque oocytes supplemented with r-hGH and demonstrated that synthesis in cumulus cells during oocyte maturation seemed to be regulated by altering expression of Cyp19A1 and 3b-HSD that metabolised the direct precursors to the end products E2 and P4 (Nyholt de Prada et al., 2010). Recently, Nakamura et al. (2012) demonstrated that GH induces preantral follicle growth and differentiation with oocyte maturation and it also acts to modulate E2 and P4 production differentially through endogenous IGF-I activity in rat granulosa cells. Eventually, data on secretion of steroid hormones by COCs during IVM with eGH, as well as data on gene expression patterns in response to eGH, are essential to obtain a more complete picture of the effects of eGH on oocyte competence. In the present study, there were clear indications of eGH-R in equine cumulus and granulosa cells, as well as in the oocyte based in our immunostaining results. In cattle, granulosa cells, primordial and secondary follicles, also had strong signals when in situ hybridization and IHC were used to detect bGH-R expression during early folliculogenesis (Kolle et al., 1998, 2001). Similarly, positive immunostaining on the equine anterior pituitary in the present study was also an indicator of eGH-R having a role in the eGH action in the horse. Thus, eGH-R may mediate a positive effect when eGH is used in culture media during IVM of equine oocytes. The effect of GH on bovine oocyte maturation is exerted via GH-R located in the cumulus cells, and that this process was not mediated by IGF-I, but probably by a G-protein-coupled mechanism through cAMP signal transduction, triggering resumption of meiosis (Bevers and Izadyar, 2002). In contrast, the effect of GH on the IVM of rat oocytes was mediated by cumulus cells, and required the intermediacy of endogenously generated cellderived IGF-I (Apa et al., 1994). The GH and GH-R expression in bovine COCs suggests that GH action in the cumulus cells might be secreted by the oocyte or by the granulosa cells (Bevers and Izadyar, 2002). In humans, hGH-R was detected in the oocyte and cumulus cells during final stages of oocyte maturation and was reported to be present during early embryonic development (Menezo et al., 2003). Based on real-time RT-PCR analysis in the present study, relative gene transcription for eGH-R was signifi-

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cantly higher in cumulus cells and COCs at the start of in vitro culture. The presence of transcript and immunoreactivity in equine cumulus cells and COCs, together with the observed responses of eGH on oocyte maturation, were interpreted as a functional eGH-R. Based on the presence of eGH-R and its mRNA we inferred that COCs and cumulus cells were a target for GH, suggesting paracrine and/or autocrine regulation of eGH on oocyte maturation. In conclusion, the principal new findings in our study were as follows: (1) exogenous eGH significantly increased P4, T4 and E2 production during oocyte maturation in vitro, whereas A4 remained constant; (2) positive immunostaining for eGH-R in cumulus cells and oocytes; and (3) presence of eGH-R in cumulus cells and COCs at the start of IVM. In conclusion, the presence of eGH-R in equine ovarian follicular structures may mediate a positive effect when eGH is used in culture during equine oocyte maturation and COCs steroidogenesis in vitro. Acknowledgements The authors thank: Paul Panich (Lethbridge Research Center) for technical assistance with recovery of COCs and Frank Ventimiglia (UCD) for assistance with confocal microscopy. This research was supported, in part, from Del Amo Program Grant (University of California, Davis and Universidad Complutense de Madrid, Spain) and The Coordination for the Improvement of Higher Education Personnel/CAPES, Brazil. References Apa, R., Lanzone, A., Miceli, F., Mastrandrea, M., Caruso, A., Mancuso, S., Canipari, R., 1994. Growth hormone induces in vitro maturation of follicle- and cumulusenclosed rat oocytes. Molecular and Cellular Endocrinology 106, 207–212. Bachelot, A., Monget, P., Imbert-Bollore, P., Coshigano, K., Kopchick, J.J., Kelly, P.A., Binart, N., 2002. Growth hormone is required for ovarian follicular growth. Endocrinology 143, 4104–4112. Belin, F., Goudet, G., Duchamp, G., Gerard, N., 2000. Intrafollicular concentrations of steroids and steroidogenic enzymes in relation to follicular development in the mare. Biology of Reproduction 62, 1335–1343. Berelowitz, M., Szabo, M., Frohman, L.A., Firestone, S., Chu, L., Hintz, R.L., 1981. Somatomedin-C mediates growth hormone negative feedback by effects on both the hypothalamus and the pituitary. Science 212, 1279–1281. Bevers, M.M., Izadyar, F., 2002. Role of growth hormone and growth hormone receptor in oocyte maturation. Molecular and Cellular Endocrinology 197, 173– 178. Borski, R.J., Tsai, W., DeMott-Friberg, R., Barkan, A.L., 1996. Regulation of somatic growth and the somatotropic axis by gonadal steroids: primary effect on insulin-like growth factor I gene expression and secretion. Endocrinology 137, 3253–3259. Caillaud, M., Gerard, N., 2009. In vivo and in vitro effects of interleukin-1beta on equine oocyte maturation and on steroidogenesis and prostaglandin synthesis in granulosa and cumulus cells. Reproduction, Fertility, and Development 21, 265–273. Carneiro, G.F., Liu, I.K., Hyde, D., Anderson, G.B., Lorenzo, P.L., Ball, B.A., 2002. Quantification and distribution of equine oocyte cortical granules during meiotic maturation and after activation. Molecular Reproduction and Development 63, 451–458. Choi, Y.H., Norris, J.D., Velez, I.C., Jacobson, C.C., Hartman, D.L., Hinrichs, K., 2013. A viable foal obtained by equine somatic cell nuclear transfer using oocytes recovered from immature follicles of live mares. Theriogenology 79, 791–796. Damiani, P., Fissore, R.A., Cibelli, J.B., Long, C.R., Balise, J.J., Robl, J.M., Duby, R.T., 1996. Evaluation of developmental competence, nuclear and ooplasmic maturation of calf oocytes. Molecular Reproduction and Development 45, 521–534. Davidson, T.R., Chamberlain, C.S., Bridges, T.S., Spicer, L.J., 2002. Effect of follicle size on in vitro production of steroids and insulin-like growth factor (IGF)-I, IGF-II, and the IGF-binding proteins by equine ovarian granulosa cells. Biology of Reproduction 66, 1640–1648. de Prada, J.K., VandeVoort, C.A., 2008. Growth hormone and in vitro maturation of rhesus macaque oocytes and subsequent embryo development. Journal of Assisted Reproduction and Genetics 25, 145–158. Ducibella, T., Anderson, E., Albertini, D.F., Aalberg, J., Rangarajan, S., 1988. Quantitative studies of changes in cortical granule number and distribution in the mouse oocyte during meiotic maturation. Developmental Biology 130, 184– 197. Ducibella, T., Duffy, P., Buetow, J., 1994. Quantification and localization of cortical granules during oogenesis in the mouse. Biology of Reproduction 50, 467–473.

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