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Expression of the orexigenic peptide ghrelin in the sheep ovary
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Domestic Animal Endocrinology 36 (2009) 89–98
Expression of the orexigenic peptide ghrelin in the sheep ovary Chenguang Du a,b , Xilingaowa a , Guifang Cao a,∗ , Caiyun Wang a , Haijun Li a , Yanhong Zhao a , Siqingaowa b , Jinshan Cao a a
College of Animal Science and Animal Medicine, Inner Mongolia Agricultural University, Hohhot 010018, China b Vocational and Technical College, Inner Mongolia Agricultural University, Baotou 014109, China Received 2 July 2008; received in revised form 13 October 2008; accepted 24 October 2008
Abstract Ghrelin has been implicated in the control of cell proliferation in reproductive tissue. Here, we provide evidence that both ghrelin mRNA and protein are present in ovarian follicles. Persistent expression of ghrelin was also demonstrated in sheep ovary throughout the estrous cycle and pregnancy. In fact, the relative mRNA levels of ghrelin varied depending on the stage of the cycle, with the highest expression during the development of the corpora lutea (CL) and minimal expression in the regressing CL. A similar pattern was seen during pregnancy. Dynamic changes in the profile of ghrelin expression during the estrous cycle and throughout pregnancy suggest a precise regulation of ovarian expression of ghrelin, which could represent a potential role for ghrelin in the regulation of luteal development. In conclusion, the presence of the ghrelin signaling system within the sheep ovary especially in the oocytes opens up the possibility of a potential regulatory role of this novel molecule in reproductive function. © 2008 Elsevier Inc. All rights reserved. Keywords: Ghrelin; Sheep ovary; Follicle; Oocyte; Corpora lutea
1. Introduction Ghrelin was identified in 1999 as the natural ligand of the GH secretagogue receptor (GHS-R1a), a 7-transmembrane G protein-coupled receptor [1]. A striking feature of ghrelin is its widespread pattern of expression [2,3]. Expression of ghrelin has been demonstrated in an array of tissues and cell types including the stomach, small intestine, pancreas, lymphocytes, placenta, kidney, lung, pituitary and brain [3]. This ubiquitous pattern of expression strongly suggests that in addition to systemic actions of the gut-derived peptide, locally produced ghrelin might have paracrine/autocrine regulatory effects in different
∗ Corresponding author. Tel.: +86 471 4303140; fax: +86 471 4303140. E-mail address: [email protected] (G. Cao).
0739-7240/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.domaniend.2008.10.007
tissues [4–6]. This may be the case in various reproductive organs such as endometrium, placenta, testis and ovary [7,8]. Interestingly, a wide range of endocrine and non-endocrine tissues, including the gonads [9], possess GHS-binding sites, and many of these tissues also have significant levels of ghrelin mRNA. Expression of ghrelin mRNA and its cognate receptor have been demonstrated in rat, pig, sheep and human gonads [10–13]. Persistent expression of the ghrelin gene was also demonstrated in rat and human ovary throughout the estrous cycle, and its relative mRNA levels varied depending on the stage of cycle [10,13]. In sheep, ghrelin immunostaining also was detected in ovarian follicles at all developmental stages, mainly in the granulosa cells [12]. At the same time, strong ghrelin immunostaining was evident in the CL of the sheep ovary, similar to the findings in the rat and human CL. These data provide further evidence for a reproductive role for this relatively new hormone.
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It is unknown whether the ghrelin system is regulated in sheep ovary and it remains equivocal as to whether ghrelin is expressed in the different stages of the oocyte. The aim of this study was to characterize in detail the pattern of expression of ghrelin in the sheep ovary, especially in the oocytes, with special attention to the cellular distribution of ghrelin peptide within the ovarian tissue, as well as the influence of the estrous cycle and pregnancy on ovarian ghrelin mRNA expression levels.
2. Materials and methods 2.1. Collection of ovine ovarian tissue and isolation of oocytes, cumulus cells, complex of granulosa cells and theca cells All experiments involving animals were approved by the animal care and use committee at the institution where the experiment was conducted. A total of 40 Mongolia sheep (12–24 months of age) were used in this study from September to January. In half of the animals (n = 20), estrus was synchronized by treatment for 8 days with a CIDR device (Hamilton, New Zealand). Appearance of estrus behaviour was detected with adult rams at 24 h after CIDR withdrawal. For RNA extraction, ovarian CL were immediately snap frozen in liquid nitrogen, while the remainder of ovaries used for cell extraction were stored in ice cold saline for transport to the laboratory. Ovarian tissues representing different stages of the estrous cycle were used from: young CL (1–3 days after ovulation, n = 6); mature CL (9–11 days after ovulation, n = 7); and regressing CL (14–16 days after ovulation, n = 7). CL from very early (<6 weeks, n = 5), early (6–8 weeks, n = 5), middle (9–14 weeks, n = 5) or late (15–18 weeks, n = 5) stages of pregnancy were also used. Ovarian CLs were sampled from 20 pregnant sheep immediately after slaughter at a local abattoir. The stage of pregnancy was estimated by measuring foetal size as previously described [14]. COCs (cumulus-oocyte complexes) were collected with a surgical blade from the surface of intact healthy antral follicles 3–5 mm in diameter. COCs having an evenly granulated cytoplasm with at least four layers of unexpanded cumulus cells (CCs) were selected. Complexes of granulosa cells (GCs) and theca cells were collected using a small scoop at the same time. Denuded oocytes were generated by vortexing for 2–3 min. Any remaining CCs were mechanically removed by repeated passage through a fine bore polished glass pipette and collected separately.
2.2. Immunohistochemistry Sheep ovaries were bisected and fixed in 4% paraformaldehyde (PFA) for 6 h. Fixed tissues were embedded in paraffin and sliced into 5 m sections, which were placed on poly-l-lysine (Sigma, USA) coated slides. Sections were deparaffinized by two 30 min exposures to xylene, then hydrated by successive 5 min washes in 95% then 85% ethanol. The remaining formaldehyde was removed from each section by incubation in 10% ammonium hydroxide in ethanol for 10 min, followed by a brief wash in distilled water. Antigen retrieval was achieved by incubating in 10 mM EDTA, pH 8.0 at 96 ◦ C for 25 min. Endogenous peroxidase was quenched by incubation in hydrogen peroxide for 20 min, then the slides were rinsed twice for 5 min in distilled water and once in 0.5 M Tris buffer, pH 7.4. Tissue sections were placed in a blocking solution for 1 h at room temperature in a humidified chamber. Blocking solution consisted of 0.1 M glycine, 1% goat serum, 0.01% Triton X-100, 1% powdered non-fat dry milk, 0.5% BSA, and 0.02% sodium azide in PBS. After blocking, tissue sections were incubated with anti-human ghrelin (Phoenix Pharmaceuticals, CA) at a 1:600 dilution overnight at 4 ◦ C. Ghrelin peptide sequences are highly conserved between species [15] and specificity of the antibody used in this study for sheep ghrelin has been previously demonstrated [12]. Slides were washed in Tris buffer twice for 5 min, then incubated with horseradish peroxidase-conjugated secondary antibody (EnVision Dual Link System; DakoCytomation). Immunostaining was revealed with liquid DAB (DakoCytomation). Negative controls were performed by omitting primary antibody from the procedure. 2.3. Immunofluorescence staining A similar immunohistochemical procedure was used for assessment of the presence and pattern of sheep antral follicles by using the same antibody. Briefly, COCs (denoted denuded CCs), oocytes and GCs were fixed for 2 h in 4% PFA at 4 ◦ C and permeabilized with 0.2% Triton X-100 for 10 min. COCs, oocytes and GCs were then incubated first with anti-human ghrelin antibody for 1 h and then with goat anti-rabbit FITC-labeled secondary antibodies (Santa Cruz Biotech) for 1 h at a 1:500 dilution. After washing for 15 min in four changes of PBS, COCs, oocytes and GCs were stained with propidium iodide (0.1 mg/mL final concentration) for 1 h, rinsed in PBS-T for 30 min, mounted on a drop of PBS containing 0.5% FBS and analyzed with a laser scanner confocal microscope (Bio-Rad MRC 1024ES) equipped
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with a Nikon (Tokyo, Japan) Diaphot inverted microscope. Simultaneously, a negative control was performed by using PBS containing 2% BSA as a substitute of the first antibody; other steps were similar to experimental groups. Each test was repeated at least four times. Staining was considered positive if the cells demonstrated green staining of the cellular cytoplasm and membrane. Red staining denoted nucleus.
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100% alcohol for 1 min for three changes, and then dipped into the absolute xylene for three changes. The slides were mounted with Eukitt mounting medium (Electron Microscopy Sciences, USA). Control sections that had no probe applied were processed in order to compare labeling between the two groups, with blue product denoting a positive signal. 2.5. RNA extraction and real-time PCR
2.4. Generation of probes and in situ hybridization Digoxigenin (DIG)-labeled RNA probes were synthesized using ghrelin cDNA generated from RNA extracted from sheep stomach as described below for CL RNA. The cDNA generated from the ghrelin primers were used for real time RT-PCR. We used 200 ng of purified PCR product (PCR Purification Kit, Qiagen, Germany) to synthesize DIG-labeled RNA probes following the manufacturer’s instructions for the DIG-RNA-Labeling Kit (Roche, Germany). Fresh ovaries were fixed in 4% PFA for 6 h, then incubated in 0.5 M sucrose overnight at 4 ◦ C. The ovaries were then embedded in paraffin, cut into 6 m sections, the sections were then hydrated in graded ethanols, washed in 0.5 × SSC for 5 min and treated for 10 min with 20 g/mL proteinase K. Following another wash with 0.5 × SSC for 10 min, 100 L of hybridization buffer was placed on each slide and the slides were prehybridized for 2 h at 42 ◦ C in a humidified chamber. Slides were incubated in 0.5 × SSC for 5 min, and the 100 L of fresh hybridization buffer containing probe was added to the slides. The slides were then incubated overnight at 55 ◦ C in a humidified chamber. Next, sections were thoroughly washed once in 2 × SSC for 15 min at room temperature, once in 1 × SSC for 5 min at RT, twice in 0.5 × SSC for 15 min at 42 ◦ C, twice in 0.1 × SSC for 15 min at 42 ◦ C, and once in maleic acid buffer (100 mM maleic acid and 150 mM NaCI, pH 7.5) for 5 min at RT. For detection of hybridization, sections were incubated with anti-digoxigenin conjugated with alkaline phosphatase (Roche Molecular Biochemicals) diluted 1:500 in 0.1 M Tris–HCl (pH 7.4), 0.1 M NaCl, with 1% blocking reagent (Roche Molecular Biochemicals). After three washes in maleic acid buffer, substrate consisting of nitroblue tetrazolium (NBT) and 5-bromocresyl-3-indolyl phosphate (BCIP) were layered over the sections. Color was allowed to develop for 2–3 h in the dark, and the reaction was stopped by dipping slides briefly in Tris–ethylenediaminetetraacetic acid buffer (10 mM Tris–HCl and 1 mM EDTA, pH 8.0). The slides were then washed with distilled water for 1 min, and then dehydrated in 95% alcohol for 1 min,
RNA was extracted from pools of 100 oocytes of remainder of 20 sheep ovaries and the associated cells (CCs, complex of GCs and theca cells) with the RNeasy Kit (Qiagen, Germany). The RNA yield from the samples were too low to be accurately quantified by spectrometry, so 6.5 L RNA aliquots were amplified. Tissue RNA was extracted from 50 mg of CL using RNAgents® Total RNA Isolation System (Promega, USA). The RNA pellets were dissolved in nuclease-free water and concentrations were measured by spectrophotometry at 260 nm. All RNA per sample was also incubated with RNase-free DNase I to remove any remaining genomic contamination. For amplification of the targets, RT and PCR were run in two separate steps, RNA was reverse transcribed with PrimeScriptTM RTase (TaKaRa, Inc. Dalian, China) following the manufacturer’s directions and performed at 37 ◦ C for 15 min for reverse transcription and then 84 ◦ C for 5 s to inactivate the reverse transcriptase. Primers used in this study were as follows: ghrelin (233 bp, GenBank accession no. AB060699) 5’-GCAGAGAAAGGAACCTAAG-3’ (forward) and 5’-GGGTTTCTTCGGCTTCTTC-3’ (reverse); -actin (208 bp, GenBank accession no. U39357) 5’-GTCACCAACTGGGACGACA-3’ (forward) and 5’-AGGCGTACAGGGACAGCA -3’ (reverse). The primers used in this study were chosen to avoid the amplification of genomic DNA. Sense oligonucleotide primer was directed toward a selected region spanning the junction of exons 1 and 2 of the gene encoding ghrelin; antisense primer was directed toward a selected region that spans the junction of exons 3 and 4 of the gene encoding ghrelin. -Actin primers also span an intron [16,17]. The threshold cycle (Ct) value represents the cycle number at which sample fluorescence rises statistically above background. Relative levels of ghrelin mRNA were quantified using SYBR® Premix Ex TaqTM (TaKaRa, Inc. Dalian, China) and a DNA Engine Opticon2 Two-Color RealTime PCR Detection System (Bio-Rad laboratories, Inc. USA). Reactions were also performed with positive (sheep stomach) and negative (water replacing cDNA)
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controls. The PCR reaction mixture (20 L) contained 10 L 2 × SYBR Green mix and 0.4 L (0.5 mM) each of forward and reverse specific primers, 2 L of cDNA and 7.2 L H2 O. The same dilution was used for both ghrelin and -actin. The PCR protocol used a denaturation step (95 ◦ C for 30 s), followed by an amplification and quantification program repeated 45 times (95 ◦ C for 20 s, 58 ◦ C for 20 s, 72 ◦ C for 30 s with a single fluorescence measurement), a melting curve program (72–95 ◦ C, with a heating rate of 0.5 ◦ C/s and continuous fluorescence measurement). Fluorescence data was acquired after the extension step during PCR reactions that contained SYBR Green. Thereafter, PCR products were analyzed by generating a melting curve. The melting curve of a product is sequence-specific and can be used to distinguish non-specific from specific PCR products. Real time PCR efficiencies of ghrelin and -actin were acquired by amplification of a dilution series of PCR products and were close to 1. Gene expression was presented using a modification of the 2−CT method, first described by Livak in PE Biosystems Sequence Detector User Bulletin 2 [18]. Expression levels of ghrelin was calculated as relative values using the 2−CT method [19], where CT = CT of the ghrelin-CT of the -actin. The sizes of RT-PCR products were confirmed by gel electrophoresis on 2% agarose gels stained with ethidium bromide, and bands were visualized by exposure to ultraviolet light. Sequences were confirmed by
Sangon Biological Engineering Technology and Services Co., Ltd. (Shanghai, China). 2.6. Statistical analysis All relative mRNA levels in oocytes, CCs, complex of GCs and theca cells, the different stages of the estrous cycle and pregnancy CLs were expressed as the mean ± S.E.M. Statistical significance was determined by one-way ANOVA with SPSS13.0 for Windows (Chicago, USA). The least significant difference (LSD) tests were applied for post hoc multiple comparison test. Differences were considered significant when P < 0.05. 3. Results 3.1. Localization of ghrelin mRNA and protein in sheep ovaries The cellular expression of ghrelin in the ovarian follicles was evaluated by in situ hybridization and immunohistochemistry. Widespread expression of ghrelin protein was observed in sheep ovary, with a detectable specific signal in oocytes as well as in somatic follicular cells. Concerning the follicular compartment, oocytes showed cytoplasmic immunostaining that was stronger in pre-antral follicles (Fig. 1A–C) and fainter in antral follicles (Fig. 1D). In addition, ghrelin
Fig. 1. Immunolocalization of ghrelin protein in sheep ovarian follicles. (A–C) Ghrelin immunostaining in oocyte and weak immunolabeling in granulosa cells of primordial follicles, primary follicles and secondary follicle, respectively. Also, with strong immunostaining in OSE (original magnification, 200×) and (D) antral follicles with strong immunostaining in both oocyte and cells are presented, cumulus and granulosa cells presented no immunolabeling (original magnification, 40×). In each panel, the insert shows the negative control. (a–d) The bottom panel is a corresponding magnification micrograph showing positive immunostaining in oocyte(Original magnification, 400×).
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Fig. 2. Laser scanning confocal microscopy images of ghrelin protein in sheep oocytes, cumulus cells and granulosa cells. (a) Negative control of cumulus cells, (b–d) ghrelin protein (green), (a’–d’) nuclei (red), (A–D) ghrelin (green) and nuclei (red) merged images. (a, a’, A) Cumulus cells, (b, b’, B) oocytes, (c, c’, C) cumulus cells, (d, d’, D) GC. The magnification was 40×, with a zoom of 1.5. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
protein was detected in oocytes of pre-antral follicles (Fig. 1a–d), with an expression profile that was roughly parallel to follicle development. Meanwhile, follicles showed variable ghrelin immunoreactivity. In detail, GCs in primordial, primary and secondary follicles
were faintly immunostained (Fig. 1, a–c). In contrast, with continued follicular development, antral follicles showed variable expression (Fig. 1d). Immunostaining was negligible in CCs and GCs but stronger in theca cells. Also, the ghrelin signal was more intense in the
Fig. 3. Localization of ghrelin mRNA in sheep ovary. Photomicrographs of sheep ovarian sections following in situ hybridization of DIG-labeled RNA probes to ghrelin mRNA. Ovarian follicles showed a wide distribution of hybridization. (A) Ghrelin mRNA is found in the oocyte, CC, GC and theca cells of the antral follicles as well as the OSE; (B) high magnification micrograph showing positive immunostaining for ghrelin in the oocyte, CC and GC; (C) ghrelin mRNA is found in secondary follicles (SF) and the GC. Scale bar, 40 m. In each panel, the insert shows the negative control.
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ovarian surface epithelium (OSE) (Fig. 1A–C). Overall, in the sheep ovary, ghrelin was immuno-localized to ovarian follicles at all developmental stages (primordial, primary, secondary and antral). In situ hybridization for ghrelin mRNA also showed a wide pattern of hybridization within the ovarian follicles. Ghrelin mRNA was clearly localized to the oocytes, CCs, GCs and theca cells, as well as the OSE (Fig. 2A and B). The presence of ghrelin mRNA was also localized to follicles at all developmental stages (primary, secondary and antral) (Fig. 2C). 3.2. Ghrelin expression in sheep ovarian antral follicles Sensitive immunofluorescence assays were used to confirm ghrelin expression in oocytes, CCs and GCs. Immunofluorescence staining confirmed immunohistochemistry results which showed that ghrelin existed in the oocytes and further identified the expression of ghrelin in CCs and GCs (Fig. 3). Ghrelin mRNA expression was evaluated in antral follicles by means of relative quantitative real time RTPCR. Real time RT-PCR assays using ghrelin-specific primers resulted in the generation of 233 bp amplicons, consistent with the known size of ghrelin mRNA. Real time RT-PCR analyses revealed that ghrelin mRNA transcripts are expressed in oocytes, CCs and GCs. Similar signals were amplified from the stomach and used as a positive control. In each sample, parallel amplification using a specific primer set for -actin served as an internal control. No products were detected from the negative controls. Relative expression levels of ghrelin did not differ between oocytes, CCs of COCs and GCs of antral follicles (Fig. 4). 3.3. Ghrelin expression in CL of the cyclic and pregnant sheep ovary Widespread expression of ghrelin mRNA and protein was observed in CL collected during the estrous cycle and pregnancy. Ghrelin mRNA and protein were clearly detected in all developmental stages of the CL, with detectable signal in luteal cells from young, mature, old, and regressing CL of the cyclic sheep ovary. Also, ghrelin mRNA and protein were detected clearly in CL from very early, early, middle or late stages of pregnancy. Ghrelin protein and mRNA signal were observed in the cytoplasm of both the larger luteal cells and small luteal cells (Fig. 5). A real-time PCR procedure was used to assess the presence of ghrelin mRNA in the CL throughout the
Fig. 4. Ghrelin mRNA in sheep oocytes, cumulus cells and complex of granulosa cells and theca cells. The top panel illustrates results from real-time PCR where ghrelin and B-actin were amplified simultaneously in each reaction. -actin was amplified in each PCR as an internal control. cDNA from a sheep stomach was used as the positive control, and samples run without template cDNA are used as the negative control. The expected 233 bp ghrelin product is detected in every sample. The bottom panel is a summary of the relative ghrelin gene expression determined by relative real-time RT-PCR. The data represent the mean ± S.E.M. (n = 5 for each group).
estrous cycle. Persistent expression of ghrelin was found in the CL throughout the estrous cycle and pregnancy. RT-PCR assays also resulted in the generation of 233 bp amplicons (Fig. 6A and B), consistent with the known size of ghrelin mRNA. Relative mRNA levels of ghrelin varied depending on the stage of the cycle, with the lowest expression levels in regressing CL and highest expression at the young and mature phases. (P < 0.05) (Fig. 6A). In the CL of pregnant sheep, gherlin expression is relatively high during the very early, early, and middle stages of pregnancy, but is lower in later pregnancy (P < 0.05) (Fig. 6B). 4. Discussion In this paper, we have characterized the cellular distribution of ghrelin mRNA and protein in sheep ovaries, and evaluated the influence of the estrous cycle and pregnancy on ghrelin mRNA expression. Our present results extend previous observations on the expression of ghrelin in sheep ovary and provide novel evidence for the presence of ghrelin mRNA and protein at all stages of follicular development (primordial, secoindary and antral) in the sheep, especially in the oocytes. More relevant for the purpose of this study, our results also provide the first evidence of dynamic
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Fig. 5. Ghrelin protein and mRNA in sheep CL from estrous cycle and pregnancy. The representative figures were chosen from mature CL of the estrous cycle and CL during the middle stage of pregnancy. Micrograph protein and mRNA positive signals in the large (LLC) and small luteal cells (SLC) for ghrelin in the CL. (A) Immunolocalization of ghrelin protein in sheep CL from midluteal phase of the estrous cycle, (B) immunolocalization of ghrelin protein in sheep CL from the middle stage of pregnancy. Original magnification, 400×. (C) Localization of ghrelin mRNA in sheep CL from midluteal phase of the estrous cycle, (D) localization of ghrelin mRNA in sheep CL from the middle stage of pregnancy. Scale bar, 40 m. In each panel, the insert shows the negative control.
changes in the profile of ghrelin expression during the CL of the estrous cycle and throughout pregnancy. Emerging evidence strongly indicates that the ghrelin signal is present in the mammalian and non-mammalian ovary. For example, ghrelin was found to be expressed in human, rat, pig, sheep and chicken ovary [10–13,20], but all above findings showed an absence of ghrelin immunoreactivity in the oocytes. In this present study, immunolocalization of ghrelin within sheep ovarian tissue revealed a wide distribution of both mRNA and the protein, with a strong signal detected in pre-antral follicles. Similar with the expression pattern in the pre-antral follicle, ghrelin protein was found in the oocyte and theca cells, but the CCs and GCs in the antral follicle presented a negligible ghrelin signal. This is inconsistent with the findings of Miller et al. [12] where there was some positive staining for ghrelin on the sheep large oocyte. It should be noted, however, that this finding was not consistent across all oocytes in their study. The
reason for such a divergence is unclear, but our results strongly indicate that oocytes are also a significant source of ghrelin. One of the interesting findings of the current study was that the ghrelin signal within the pre-antral and antral follicles showed different levels of expression. To note, the reason that CCs and GCs of antral follicles showed negative results may be due to levels of ghrelin in GCs that are below the sensitivity threshold of our immunohistochemical assays. This is why immunofluorescence staining was applied to detect the expression of ghrelin in the antral follicles. The results indicated that the different cell types all showed an intense positive signal in the antral follicle. Our results supported the previous view that ghrelin is expressed in the GCs [12], but the fluctuations in protein expression in the different cell types of the antral follicles may be related to the follicle development. However, there were no differences observed at the mRNA level in the antral follicles using in situ
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Fig. 6. Ghrelin mRNA in sheep CL from the different developmetnat stages of the estrous cycle and pregnancy. The top panel illustrates results from real-time RT-PCR where ghrelin and -actin were amplified simultaneously in each reaction from the different developmental stages of the sheep CL. cDNA from a sheep stomach was used as the positive control, and samples run without template cDNA are used as the negative control. The expected 233 bp ghrelin product is detected in every sample. The bottom panel is a summary of the relative ghrelin gene expression determined by relative real-time RT-PCR. The data represent the mean ± S.E.M. (n = 5 for each group). (A) Ghrelin mRNA in sheep CL from young, mature and regressing CL. (B) mRNA was extracted from very early, early, middle, and late pregnancy CL.
hybridization and real time PCR. Hence, the relationship between mRNA and protein must be investigated in future studies. Interestingly, as the follicles developed, ghrelin staining decreased in the oocytes of antral follicles. Oocyte control of folliculogenesis is important in reproduction because perturbed expression of oocyte factors has profound effects on maturation and fertility [21]. Therefore, ghrelin may be influencing growth of the oocyte by affecting cellular proliferation. The ability of ghrelin to modulate cell proliferation has been observed in adipocyte, cardiomyocyte, osteoblast and pituitary cell lines [4,6], as well as in breast, lung and thyroid carcinoma cell lines [5,22,23]. In fact, in a rat pituitary somatotroph cell line, ghrelin exerts a proliferative effect via the mitogen-activated protein kinase (MAPK) pathway [24]. The pre-antral to antral follicle transition marks an important developmental step during follicular development where a subset of the pre-antral granulosa cells, probably those directly associated with the oocyte, differentiate into cumulus cells with competence to undergo cumulus expansion after the preovulatory LH surge [25]. GnRH and LH are important regulators for folliculogenesis, luteinization, and establishment of pregnancy as well [26,27]. Hence, the roles of ghrelin in follicular development and survival may depend on gonadotropin status and developmental state of the follicle.
During the ovulatory cycle, the OSE is subject to a series of injury and repair processes associated with follicular rupture and CL formation, which involve natural inflammatory events [28]. Expression of ghrelin within the OSE, which we have confirmed here, supports the theory that ghrelin play a vital role in OSE cells [12]. OSE is an obligate component of the ovulatory process as it plays an essential role during follicle rupture at the ovarian surface and formation of CL. Following ovulation, the small luteal cells of the CL are thought to originate from the follicular theca cells, whereas the large luteal cells are of GC origin. To ascertain the physiological regulation of ghrelin gene expression in sheep ovarian CL, relative mRNA levels were assessed during the different stages of the estrous cycle and pregnancy. Interestingly, ghrelin mRNA levels varied significantly depending on the phase of the estrous cycle. Ghrelin mRNA levels were highest at the beginning of the formation and mature phases, and lower during regression. Notably, a dynamic pattern of ovarian ghrelin mRNA expression was described in rat showing that ghrelin mRNA level varied depending on the phase of the cycle, with the lowest expression level in the proestrus and maximal value in the diestrus phase [10]. In addition, ovarian CL ghrelin gene expression was monitored throughout pregnancy. In our analysis, expression levels of ghrelin were higher from very early-pregnancy to mid-pregnancy, and significantly decreased later in gestation. Most studies on the
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biochemistry and structure of the CL suggest that during late pregnancy the CL of the rat is at the very early stages of structural regression, with no changes at the morphological level, but with changes at the molecular level (functional regression). Our results supported this report despite differences in animal species. In conclusion, we have demonstrated that the peptide ghrelin is expressed in sheep ovarian follicles at all developmental stages as well as in functional CL. Dynamic changes in the profile of ovarian expression of ghrelin mRNA during the estrous cycle and pregnancy are highly suggestive of a finely tuned regulatory network. Overall, the presence of the ghrelin signaling system within the sheep ovary especially in the oocytes opens up the possibility of a potential regulatory role of this novel molecule in reproductive function. Acknowledgements This work was supported by grants from National Natural Science Foundation of China (30660128 and 30860201) and Natural Science Foundation of Inner Mongolia (200711020506). The authors thank Mr. Yongmiao Feng and Dr. Bo Tang for their assistance and technical support during the studies. We also thank Dr. Hua Shao for his help with revision of the manuscript. References [1] Kojima M, Hosoda H, Date Y, Nakazato M, Matsuo H, Kangawa K. Ghrelin is a growth-hormone-releasing acylated peptide from stomach. Nature 1999;402:656–60. [2] van der Lely AJ, Tschop M, Heiman ML, Ghigo E. Biological, physiological, pathophysiological, and pharmacological aspects of ghrelin. Endocr Rev 2004;25:426–57. [3] Gualillo O, Lago F, Gomez-Reino J, Casanueva FF, Dieguez C. Ghrelin, a widespread hormone: insights into molecular and cellular regulation of its expression and mechanism of action. FEBS Lett 2003;552:105–9. [4] Duxbury MS, Waseem T, Ito H, Robinson MK, Zinner MJ, Ashley SW, et al. Ghrelin promotes pancreatic adenocarcinoma cellular proliferation and invasiveness. Biochem Biophys Res Commun 2003;309:464–8. [5] Xia Q, Pang W, Pan H, Zheng Y, Kang JS, Zhu SG. Effects of ghrelin on the proliferation and secretion of splenic T lymphocytes in mice. Regul Pept 2004;122:173–8. [6] Maccarinelli G, Sibilia V, Torsello A, Raimondo F, Pitto M, Giustina A, et al. Ghrelin regulates proliferation and differentiation of osteoblastic cells. J Endocrinol 2005;184:249–56. [7] Garcia MC, Lopez M, Alvarez CV, Casanueva F, Tena-Sempere M, Dieguez C. Role of ghrelin in reproduction. Reproduction 2007;133:531–40. [8] Harrison JL, Adam CL, Brown YA, Wallace JM, Aitken RP, Lea RG, et al. An immunohistochemical study of the localization and developmental expression of ghrelin and its functional receptor in the ovine placenta. Reprod Biol Endocrinol 2007;5:25.
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[9] Tena-Sempere M. Ghrelin and reproduction: ghrelin as novel regulator of the gonadotropic axis. Vitam Horm 2008;77:285–90. [10] Caminos JE, Tena-Sempere M, Gaytan F, Sanchez-Criado JE, Barreiro ML, Nogueiras R, et al. Expression of ghrelin in the cyclic and pregnant rat ovary. Endocrinology 2003;144:1594– 602. [11] Zhang W, Lei Z, Su J, Chen S. Expression of ghrelin in the porcine hypothalamo-pituitary–ovary axis during the estrous cycle. Anim Reprod Sci 2008;109:356–67. [12] Miller DW, Harrison JL, Brown YA, Doyle U, Lindsay A, Adam CL, et al. Immunohistochemical evidence for an endocrine/ paracrine role for ghrelin in the reproductive tissues of sheep. Reprod Biol Endocrinol 2005;3:60. [13] Gaytan F, Barreiro ML, Chopin LK, Herington AC, Morales C, Pinilla L, et al. Immunolocalization of ghrelin and its functional receptor, the type 1a growth hormone secretagogue receptor, in the cyclic human ovary. J Clin Endocrinol Metab 2003;88:879– 87. [14] Sivachelvan MN, Ali MG, Chibuzo GA. Foetal age estimation in sheep and goats. Small Ruminant Res 1996;19:69–76. [15] Lazarczyk MA, Lazarczyk M, Grzela T. Ghrelin: a recently discovered gut-brain peptide (review). Int J Mol Med 2003;12:279–87. [16] Jeong YJ, Choi HW, Shin HS, Cui XS, Kim NH, Gerton GL, et al. Optimization of real time RT-PCR methods for the analysis of gene expression in mouse eggs and preimplantation embryos. Mol Reprod Dev 2005;71:284–9. [17] Succu S, Bebbere D, Bogliolo L, Ariu F, Fois S, Leoni GG, et al. Vitrification of in vitro matured ovine oocytes affects in vitro preimplantation development and mRNA abundance. Mol Reprod Dev 2008;75:538–46. [18] Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods 2001;25:402–8. [19] Tropea A, Tiberi F, Minici F, Orlando M, Gangale MF, Romani F, et al. Ghrelin affects the release of luteolytic and luteotropic factors in human luteal cells. J Clin Endocrinol Metab 2007;92:3239–45. [20] Sirotkin AV, Grossmann R, Maria-Peon MT, Roa J, Tena-Sempere M, Klein S. Novel expression and functional role of ghrelin in chicken ovary. Mol Cell Endocrinol 2006;257–258:15–25. [21] Pangas SA, Rajkovic A. Transcriptional regulation of early oogenesis: in search of masters. Hum Reprod Update 2006;12:65– 76. [22] Kim SW, Her SJ, Park SJ, Kim D, Park KS, Lee HK, et al. Ghrelin stimulates proliferation and differentiation and inhibits apoptosis in osteoblastic MC3T3-E1 cells. Bone 2005;37:359–69. [23] Groschl M, Topf HG, Bohlender J, Zenk J, Klussmann S, Dotsch J, et al. Identification of ghrelin in human saliva: production by the salivary glands and potential role in proliferation of oral keratinocytes. Clin Chem 2005;51:997–1006. [24] Nanzer AM, Khalaf S, Mozid AM, Fowkes RC, Patel MV, Burrin JM, et al. Ghrelin exerts a proliferative effect on a rat pituitary somatotroph cell line via the mitogen-activated protein kinase pathway. Eur J Endocrinol 2004;151:233–40. [25] Diaz FJ, Wigglesworth K, Eppig JJ. Oocytes are required for the preantral granulosa cell to cumulus cell transition in mice. Dev Biol 2007;305:300–11. [26] Fernandez-Fernandez R, Martini AC, Navarro VM, Castellano JM, Dieguez C, Aguilar E, et al. Novel signals for the integration of energy balance and reproduction. Mol Cell Endocrinol 2006;254–255:127–32.
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[27] Filicori M, Cognigni GE, Taraborrelli S, Spettoli D, Ciampaglia W, de Tabarelli F, et al. Luteinzing hormone activity in menotropins optimizes folliculogenesis and treatment in controlled ovarian stimulation. J Clin Endocrinol Metab 2001;86:337–43.
[28] Gaytan F, Morales C, Barreiro ML, Jeffery P, Chopin LK, Herington AC, et al. Expression of growth hormone secretagogue receptor type 1a, the functional ghrelin receptor, in human ovarian surface epithelium, mullerian duct derivatives, and ovarian tumors. J Clin Endocrinol Metab 2005;90:1798–804.
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Expression of the orexigenic peptide ghrelin in the sheep ovary Chenguang Du a,b , Xilingaowa a , Guifang Cao a,∗ , Caiyun Wang a , Haijun Li a , Yanhong Zhao a , Siqingaowa b , Jinshan Cao a a
College of Animal Science and Animal Medicine, Inner Mongolia Agricultural University, Hohhot 010018, China b Vocational and Technical College, Inner Mongolia Agricultural University, Baotou 014109, China Received 2 July 2008; received in revised form 13 October 2008; accepted 24 October 2008
Abstract Ghrelin has been implicated in the control of cell proliferation in reproductive tissue. Here, we provide evidence that both ghrelin mRNA and protein are present in ovarian follicles. Persistent expression of ghrelin was also demonstrated in sheep ovary throughout the estrous cycle and pregnancy. In fact, the relative mRNA levels of ghrelin varied depending on the stage of the cycle, with the highest expression during the development of the corpora lutea (CL) and minimal expression in the regressing CL. A similar pattern was seen during pregnancy. Dynamic changes in the profile of ghrelin expression during the estrous cycle and throughout pregnancy suggest a precise regulation of ovarian expression of ghrelin, which could represent a potential role for ghrelin in the regulation of luteal development. In conclusion, the presence of the ghrelin signaling system within the sheep ovary especially in the oocytes opens up the possibility of a potential regulatory role of this novel molecule in reproductive function. © 2008 Elsevier Inc. All rights reserved. Keywords: Ghrelin; Sheep ovary; Follicle; Oocyte; Corpora lutea
1. Introduction Ghrelin was identified in 1999 as the natural ligand of the GH secretagogue receptor (GHS-R1a), a 7-transmembrane G protein-coupled receptor [1]. A striking feature of ghrelin is its widespread pattern of expression [2,3]. Expression of ghrelin has been demonstrated in an array of tissues and cell types including the stomach, small intestine, pancreas, lymphocytes, placenta, kidney, lung, pituitary and brain [3]. This ubiquitous pattern of expression strongly suggests that in addition to systemic actions of the gut-derived peptide, locally produced ghrelin might have paracrine/autocrine regulatory effects in different
∗ Corresponding author. Tel.: +86 471 4303140; fax: +86 471 4303140. E-mail address: [email protected] (G. Cao).
0739-7240/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.domaniend.2008.10.007
tissues [4–6]. This may be the case in various reproductive organs such as endometrium, placenta, testis and ovary [7,8]. Interestingly, a wide range of endocrine and non-endocrine tissues, including the gonads [9], possess GHS-binding sites, and many of these tissues also have significant levels of ghrelin mRNA. Expression of ghrelin mRNA and its cognate receptor have been demonstrated in rat, pig, sheep and human gonads [10–13]. Persistent expression of the ghrelin gene was also demonstrated in rat and human ovary throughout the estrous cycle, and its relative mRNA levels varied depending on the stage of cycle [10,13]. In sheep, ghrelin immunostaining also was detected in ovarian follicles at all developmental stages, mainly in the granulosa cells [12]. At the same time, strong ghrelin immunostaining was evident in the CL of the sheep ovary, similar to the findings in the rat and human CL. These data provide further evidence for a reproductive role for this relatively new hormone.
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It is unknown whether the ghrelin system is regulated in sheep ovary and it remains equivocal as to whether ghrelin is expressed in the different stages of the oocyte. The aim of this study was to characterize in detail the pattern of expression of ghrelin in the sheep ovary, especially in the oocytes, with special attention to the cellular distribution of ghrelin peptide within the ovarian tissue, as well as the influence of the estrous cycle and pregnancy on ovarian ghrelin mRNA expression levels.
2. Materials and methods 2.1. Collection of ovine ovarian tissue and isolation of oocytes, cumulus cells, complex of granulosa cells and theca cells All experiments involving animals were approved by the animal care and use committee at the institution where the experiment was conducted. A total of 40 Mongolia sheep (12–24 months of age) were used in this study from September to January. In half of the animals (n = 20), estrus was synchronized by treatment for 8 days with a CIDR device (Hamilton, New Zealand). Appearance of estrus behaviour was detected with adult rams at 24 h after CIDR withdrawal. For RNA extraction, ovarian CL were immediately snap frozen in liquid nitrogen, while the remainder of ovaries used for cell extraction were stored in ice cold saline for transport to the laboratory. Ovarian tissues representing different stages of the estrous cycle were used from: young CL (1–3 days after ovulation, n = 6); mature CL (9–11 days after ovulation, n = 7); and regressing CL (14–16 days after ovulation, n = 7). CL from very early (<6 weeks, n = 5), early (6–8 weeks, n = 5), middle (9–14 weeks, n = 5) or late (15–18 weeks, n = 5) stages of pregnancy were also used. Ovarian CLs were sampled from 20 pregnant sheep immediately after slaughter at a local abattoir. The stage of pregnancy was estimated by measuring foetal size as previously described [14]. COCs (cumulus-oocyte complexes) were collected with a surgical blade from the surface of intact healthy antral follicles 3–5 mm in diameter. COCs having an evenly granulated cytoplasm with at least four layers of unexpanded cumulus cells (CCs) were selected. Complexes of granulosa cells (GCs) and theca cells were collected using a small scoop at the same time. Denuded oocytes were generated by vortexing for 2–3 min. Any remaining CCs were mechanically removed by repeated passage through a fine bore polished glass pipette and collected separately.
2.2. Immunohistochemistry Sheep ovaries were bisected and fixed in 4% paraformaldehyde (PFA) for 6 h. Fixed tissues were embedded in paraffin and sliced into 5 m sections, which were placed on poly-l-lysine (Sigma, USA) coated slides. Sections were deparaffinized by two 30 min exposures to xylene, then hydrated by successive 5 min washes in 95% then 85% ethanol. The remaining formaldehyde was removed from each section by incubation in 10% ammonium hydroxide in ethanol for 10 min, followed by a brief wash in distilled water. Antigen retrieval was achieved by incubating in 10 mM EDTA, pH 8.0 at 96 ◦ C for 25 min. Endogenous peroxidase was quenched by incubation in hydrogen peroxide for 20 min, then the slides were rinsed twice for 5 min in distilled water and once in 0.5 M Tris buffer, pH 7.4. Tissue sections were placed in a blocking solution for 1 h at room temperature in a humidified chamber. Blocking solution consisted of 0.1 M glycine, 1% goat serum, 0.01% Triton X-100, 1% powdered non-fat dry milk, 0.5% BSA, and 0.02% sodium azide in PBS. After blocking, tissue sections were incubated with anti-human ghrelin (Phoenix Pharmaceuticals, CA) at a 1:600 dilution overnight at 4 ◦ C. Ghrelin peptide sequences are highly conserved between species [15] and specificity of the antibody used in this study for sheep ghrelin has been previously demonstrated [12]. Slides were washed in Tris buffer twice for 5 min, then incubated with horseradish peroxidase-conjugated secondary antibody (EnVision Dual Link System; DakoCytomation). Immunostaining was revealed with liquid DAB (DakoCytomation). Negative controls were performed by omitting primary antibody from the procedure. 2.3. Immunofluorescence staining A similar immunohistochemical procedure was used for assessment of the presence and pattern of sheep antral follicles by using the same antibody. Briefly, COCs (denoted denuded CCs), oocytes and GCs were fixed for 2 h in 4% PFA at 4 ◦ C and permeabilized with 0.2% Triton X-100 for 10 min. COCs, oocytes and GCs were then incubated first with anti-human ghrelin antibody for 1 h and then with goat anti-rabbit FITC-labeled secondary antibodies (Santa Cruz Biotech) for 1 h at a 1:500 dilution. After washing for 15 min in four changes of PBS, COCs, oocytes and GCs were stained with propidium iodide (0.1 mg/mL final concentration) for 1 h, rinsed in PBS-T for 30 min, mounted on a drop of PBS containing 0.5% FBS and analyzed with a laser scanner confocal microscope (Bio-Rad MRC 1024ES) equipped
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with a Nikon (Tokyo, Japan) Diaphot inverted microscope. Simultaneously, a negative control was performed by using PBS containing 2% BSA as a substitute of the first antibody; other steps were similar to experimental groups. Each test was repeated at least four times. Staining was considered positive if the cells demonstrated green staining of the cellular cytoplasm and membrane. Red staining denoted nucleus.
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100% alcohol for 1 min for three changes, and then dipped into the absolute xylene for three changes. The slides were mounted with Eukitt mounting medium (Electron Microscopy Sciences, USA). Control sections that had no probe applied were processed in order to compare labeling between the two groups, with blue product denoting a positive signal. 2.5. RNA extraction and real-time PCR
2.4. Generation of probes and in situ hybridization Digoxigenin (DIG)-labeled RNA probes were synthesized using ghrelin cDNA generated from RNA extracted from sheep stomach as described below for CL RNA. The cDNA generated from the ghrelin primers were used for real time RT-PCR. We used 200 ng of purified PCR product (PCR Purification Kit, Qiagen, Germany) to synthesize DIG-labeled RNA probes following the manufacturer’s instructions for the DIG-RNA-Labeling Kit (Roche, Germany). Fresh ovaries were fixed in 4% PFA for 6 h, then incubated in 0.5 M sucrose overnight at 4 ◦ C. The ovaries were then embedded in paraffin, cut into 6 m sections, the sections were then hydrated in graded ethanols, washed in 0.5 × SSC for 5 min and treated for 10 min with 20 g/mL proteinase K. Following another wash with 0.5 × SSC for 10 min, 100 L of hybridization buffer was placed on each slide and the slides were prehybridized for 2 h at 42 ◦ C in a humidified chamber. Slides were incubated in 0.5 × SSC for 5 min, and the 100 L of fresh hybridization buffer containing probe was added to the slides. The slides were then incubated overnight at 55 ◦ C in a humidified chamber. Next, sections were thoroughly washed once in 2 × SSC for 15 min at room temperature, once in 1 × SSC for 5 min at RT, twice in 0.5 × SSC for 15 min at 42 ◦ C, twice in 0.1 × SSC for 15 min at 42 ◦ C, and once in maleic acid buffer (100 mM maleic acid and 150 mM NaCI, pH 7.5) for 5 min at RT. For detection of hybridization, sections were incubated with anti-digoxigenin conjugated with alkaline phosphatase (Roche Molecular Biochemicals) diluted 1:500 in 0.1 M Tris–HCl (pH 7.4), 0.1 M NaCl, with 1% blocking reagent (Roche Molecular Biochemicals). After three washes in maleic acid buffer, substrate consisting of nitroblue tetrazolium (NBT) and 5-bromocresyl-3-indolyl phosphate (BCIP) were layered over the sections. Color was allowed to develop for 2–3 h in the dark, and the reaction was stopped by dipping slides briefly in Tris–ethylenediaminetetraacetic acid buffer (10 mM Tris–HCl and 1 mM EDTA, pH 8.0). The slides were then washed with distilled water for 1 min, and then dehydrated in 95% alcohol for 1 min,
RNA was extracted from pools of 100 oocytes of remainder of 20 sheep ovaries and the associated cells (CCs, complex of GCs and theca cells) with the RNeasy Kit (Qiagen, Germany). The RNA yield from the samples were too low to be accurately quantified by spectrometry, so 6.5 L RNA aliquots were amplified. Tissue RNA was extracted from 50 mg of CL using RNAgents® Total RNA Isolation System (Promega, USA). The RNA pellets were dissolved in nuclease-free water and concentrations were measured by spectrophotometry at 260 nm. All RNA per sample was also incubated with RNase-free DNase I to remove any remaining genomic contamination. For amplification of the targets, RT and PCR were run in two separate steps, RNA was reverse transcribed with PrimeScriptTM RTase (TaKaRa, Inc. Dalian, China) following the manufacturer’s directions and performed at 37 ◦ C for 15 min for reverse transcription and then 84 ◦ C for 5 s to inactivate the reverse transcriptase. Primers used in this study were as follows: ghrelin (233 bp, GenBank accession no. AB060699) 5’-GCAGAGAAAGGAACCTAAG-3’ (forward) and 5’-GGGTTTCTTCGGCTTCTTC-3’ (reverse); -actin (208 bp, GenBank accession no. U39357) 5’-GTCACCAACTGGGACGACA-3’ (forward) and 5’-AGGCGTACAGGGACAGCA -3’ (reverse). The primers used in this study were chosen to avoid the amplification of genomic DNA. Sense oligonucleotide primer was directed toward a selected region spanning the junction of exons 1 and 2 of the gene encoding ghrelin; antisense primer was directed toward a selected region that spans the junction of exons 3 and 4 of the gene encoding ghrelin. -Actin primers also span an intron [16,17]. The threshold cycle (Ct) value represents the cycle number at which sample fluorescence rises statistically above background. Relative levels of ghrelin mRNA were quantified using SYBR® Premix Ex TaqTM (TaKaRa, Inc. Dalian, China) and a DNA Engine Opticon2 Two-Color RealTime PCR Detection System (Bio-Rad laboratories, Inc. USA). Reactions were also performed with positive (sheep stomach) and negative (water replacing cDNA)
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controls. The PCR reaction mixture (20 L) contained 10 L 2 × SYBR Green mix and 0.4 L (0.5 mM) each of forward and reverse specific primers, 2 L of cDNA and 7.2 L H2 O. The same dilution was used for both ghrelin and -actin. The PCR protocol used a denaturation step (95 ◦ C for 30 s), followed by an amplification and quantification program repeated 45 times (95 ◦ C for 20 s, 58 ◦ C for 20 s, 72 ◦ C for 30 s with a single fluorescence measurement), a melting curve program (72–95 ◦ C, with a heating rate of 0.5 ◦ C/s and continuous fluorescence measurement). Fluorescence data was acquired after the extension step during PCR reactions that contained SYBR Green. Thereafter, PCR products were analyzed by generating a melting curve. The melting curve of a product is sequence-specific and can be used to distinguish non-specific from specific PCR products. Real time PCR efficiencies of ghrelin and -actin were acquired by amplification of a dilution series of PCR products and were close to 1. Gene expression was presented using a modification of the 2−CT method, first described by Livak in PE Biosystems Sequence Detector User Bulletin 2 [18]. Expression levels of ghrelin was calculated as relative values using the 2−CT method [19], where CT = CT of the ghrelin-CT of the -actin. The sizes of RT-PCR products were confirmed by gel electrophoresis on 2% agarose gels stained with ethidium bromide, and bands were visualized by exposure to ultraviolet light. Sequences were confirmed by
Sangon Biological Engineering Technology and Services Co., Ltd. (Shanghai, China). 2.6. Statistical analysis All relative mRNA levels in oocytes, CCs, complex of GCs and theca cells, the different stages of the estrous cycle and pregnancy CLs were expressed as the mean ± S.E.M. Statistical significance was determined by one-way ANOVA with SPSS13.0 for Windows (Chicago, USA). The least significant difference (LSD) tests were applied for post hoc multiple comparison test. Differences were considered significant when P < 0.05. 3. Results 3.1. Localization of ghrelin mRNA and protein in sheep ovaries The cellular expression of ghrelin in the ovarian follicles was evaluated by in situ hybridization and immunohistochemistry. Widespread expression of ghrelin protein was observed in sheep ovary, with a detectable specific signal in oocytes as well as in somatic follicular cells. Concerning the follicular compartment, oocytes showed cytoplasmic immunostaining that was stronger in pre-antral follicles (Fig. 1A–C) and fainter in antral follicles (Fig. 1D). In addition, ghrelin
Fig. 1. Immunolocalization of ghrelin protein in sheep ovarian follicles. (A–C) Ghrelin immunostaining in oocyte and weak immunolabeling in granulosa cells of primordial follicles, primary follicles and secondary follicle, respectively. Also, with strong immunostaining in OSE (original magnification, 200×) and (D) antral follicles with strong immunostaining in both oocyte and cells are presented, cumulus and granulosa cells presented no immunolabeling (original magnification, 40×). In each panel, the insert shows the negative control. (a–d) The bottom panel is a corresponding magnification micrograph showing positive immunostaining in oocyte(Original magnification, 400×).
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Fig. 2. Laser scanning confocal microscopy images of ghrelin protein in sheep oocytes, cumulus cells and granulosa cells. (a) Negative control of cumulus cells, (b–d) ghrelin protein (green), (a’–d’) nuclei (red), (A–D) ghrelin (green) and nuclei (red) merged images. (a, a’, A) Cumulus cells, (b, b’, B) oocytes, (c, c’, C) cumulus cells, (d, d’, D) GC. The magnification was 40×, with a zoom of 1.5. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
protein was detected in oocytes of pre-antral follicles (Fig. 1a–d), with an expression profile that was roughly parallel to follicle development. Meanwhile, follicles showed variable ghrelin immunoreactivity. In detail, GCs in primordial, primary and secondary follicles
were faintly immunostained (Fig. 1, a–c). In contrast, with continued follicular development, antral follicles showed variable expression (Fig. 1d). Immunostaining was negligible in CCs and GCs but stronger in theca cells. Also, the ghrelin signal was more intense in the
Fig. 3. Localization of ghrelin mRNA in sheep ovary. Photomicrographs of sheep ovarian sections following in situ hybridization of DIG-labeled RNA probes to ghrelin mRNA. Ovarian follicles showed a wide distribution of hybridization. (A) Ghrelin mRNA is found in the oocyte, CC, GC and theca cells of the antral follicles as well as the OSE; (B) high magnification micrograph showing positive immunostaining for ghrelin in the oocyte, CC and GC; (C) ghrelin mRNA is found in secondary follicles (SF) and the GC. Scale bar, 40 m. In each panel, the insert shows the negative control.
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ovarian surface epithelium (OSE) (Fig. 1A–C). Overall, in the sheep ovary, ghrelin was immuno-localized to ovarian follicles at all developmental stages (primordial, primary, secondary and antral). In situ hybridization for ghrelin mRNA also showed a wide pattern of hybridization within the ovarian follicles. Ghrelin mRNA was clearly localized to the oocytes, CCs, GCs and theca cells, as well as the OSE (Fig. 2A and B). The presence of ghrelin mRNA was also localized to follicles at all developmental stages (primary, secondary and antral) (Fig. 2C). 3.2. Ghrelin expression in sheep ovarian antral follicles Sensitive immunofluorescence assays were used to confirm ghrelin expression in oocytes, CCs and GCs. Immunofluorescence staining confirmed immunohistochemistry results which showed that ghrelin existed in the oocytes and further identified the expression of ghrelin in CCs and GCs (Fig. 3). Ghrelin mRNA expression was evaluated in antral follicles by means of relative quantitative real time RTPCR. Real time RT-PCR assays using ghrelin-specific primers resulted in the generation of 233 bp amplicons, consistent with the known size of ghrelin mRNA. Real time RT-PCR analyses revealed that ghrelin mRNA transcripts are expressed in oocytes, CCs and GCs. Similar signals were amplified from the stomach and used as a positive control. In each sample, parallel amplification using a specific primer set for -actin served as an internal control. No products were detected from the negative controls. Relative expression levels of ghrelin did not differ between oocytes, CCs of COCs and GCs of antral follicles (Fig. 4). 3.3. Ghrelin expression in CL of the cyclic and pregnant sheep ovary Widespread expression of ghrelin mRNA and protein was observed in CL collected during the estrous cycle and pregnancy. Ghrelin mRNA and protein were clearly detected in all developmental stages of the CL, with detectable signal in luteal cells from young, mature, old, and regressing CL of the cyclic sheep ovary. Also, ghrelin mRNA and protein were detected clearly in CL from very early, early, middle or late stages of pregnancy. Ghrelin protein and mRNA signal were observed in the cytoplasm of both the larger luteal cells and small luteal cells (Fig. 5). A real-time PCR procedure was used to assess the presence of ghrelin mRNA in the CL throughout the
Fig. 4. Ghrelin mRNA in sheep oocytes, cumulus cells and complex of granulosa cells and theca cells. The top panel illustrates results from real-time PCR where ghrelin and B-actin were amplified simultaneously in each reaction. -actin was amplified in each PCR as an internal control. cDNA from a sheep stomach was used as the positive control, and samples run without template cDNA are used as the negative control. The expected 233 bp ghrelin product is detected in every sample. The bottom panel is a summary of the relative ghrelin gene expression determined by relative real-time RT-PCR. The data represent the mean ± S.E.M. (n = 5 for each group).
estrous cycle. Persistent expression of ghrelin was found in the CL throughout the estrous cycle and pregnancy. RT-PCR assays also resulted in the generation of 233 bp amplicons (Fig. 6A and B), consistent with the known size of ghrelin mRNA. Relative mRNA levels of ghrelin varied depending on the stage of the cycle, with the lowest expression levels in regressing CL and highest expression at the young and mature phases. (P < 0.05) (Fig. 6A). In the CL of pregnant sheep, gherlin expression is relatively high during the very early, early, and middle stages of pregnancy, but is lower in later pregnancy (P < 0.05) (Fig. 6B). 4. Discussion In this paper, we have characterized the cellular distribution of ghrelin mRNA and protein in sheep ovaries, and evaluated the influence of the estrous cycle and pregnancy on ghrelin mRNA expression. Our present results extend previous observations on the expression of ghrelin in sheep ovary and provide novel evidence for the presence of ghrelin mRNA and protein at all stages of follicular development (primordial, secoindary and antral) in the sheep, especially in the oocytes. More relevant for the purpose of this study, our results also provide the first evidence of dynamic
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Fig. 5. Ghrelin protein and mRNA in sheep CL from estrous cycle and pregnancy. The representative figures were chosen from mature CL of the estrous cycle and CL during the middle stage of pregnancy. Micrograph protein and mRNA positive signals in the large (LLC) and small luteal cells (SLC) for ghrelin in the CL. (A) Immunolocalization of ghrelin protein in sheep CL from midluteal phase of the estrous cycle, (B) immunolocalization of ghrelin protein in sheep CL from the middle stage of pregnancy. Original magnification, 400×. (C) Localization of ghrelin mRNA in sheep CL from midluteal phase of the estrous cycle, (D) localization of ghrelin mRNA in sheep CL from the middle stage of pregnancy. Scale bar, 40 m. In each panel, the insert shows the negative control.
changes in the profile of ghrelin expression during the CL of the estrous cycle and throughout pregnancy. Emerging evidence strongly indicates that the ghrelin signal is present in the mammalian and non-mammalian ovary. For example, ghrelin was found to be expressed in human, rat, pig, sheep and chicken ovary [10–13,20], but all above findings showed an absence of ghrelin immunoreactivity in the oocytes. In this present study, immunolocalization of ghrelin within sheep ovarian tissue revealed a wide distribution of both mRNA and the protein, with a strong signal detected in pre-antral follicles. Similar with the expression pattern in the pre-antral follicle, ghrelin protein was found in the oocyte and theca cells, but the CCs and GCs in the antral follicle presented a negligible ghrelin signal. This is inconsistent with the findings of Miller et al. [12] where there was some positive staining for ghrelin on the sheep large oocyte. It should be noted, however, that this finding was not consistent across all oocytes in their study. The
reason for such a divergence is unclear, but our results strongly indicate that oocytes are also a significant source of ghrelin. One of the interesting findings of the current study was that the ghrelin signal within the pre-antral and antral follicles showed different levels of expression. To note, the reason that CCs and GCs of antral follicles showed negative results may be due to levels of ghrelin in GCs that are below the sensitivity threshold of our immunohistochemical assays. This is why immunofluorescence staining was applied to detect the expression of ghrelin in the antral follicles. The results indicated that the different cell types all showed an intense positive signal in the antral follicle. Our results supported the previous view that ghrelin is expressed in the GCs [12], but the fluctuations in protein expression in the different cell types of the antral follicles may be related to the follicle development. However, there were no differences observed at the mRNA level in the antral follicles using in situ
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Fig. 6. Ghrelin mRNA in sheep CL from the different developmetnat stages of the estrous cycle and pregnancy. The top panel illustrates results from real-time RT-PCR where ghrelin and -actin were amplified simultaneously in each reaction from the different developmental stages of the sheep CL. cDNA from a sheep stomach was used as the positive control, and samples run without template cDNA are used as the negative control. The expected 233 bp ghrelin product is detected in every sample. The bottom panel is a summary of the relative ghrelin gene expression determined by relative real-time RT-PCR. The data represent the mean ± S.E.M. (n = 5 for each group). (A) Ghrelin mRNA in sheep CL from young, mature and regressing CL. (B) mRNA was extracted from very early, early, middle, and late pregnancy CL.
hybridization and real time PCR. Hence, the relationship between mRNA and protein must be investigated in future studies. Interestingly, as the follicles developed, ghrelin staining decreased in the oocytes of antral follicles. Oocyte control of folliculogenesis is important in reproduction because perturbed expression of oocyte factors has profound effects on maturation and fertility [21]. Therefore, ghrelin may be influencing growth of the oocyte by affecting cellular proliferation. The ability of ghrelin to modulate cell proliferation has been observed in adipocyte, cardiomyocyte, osteoblast and pituitary cell lines [4,6], as well as in breast, lung and thyroid carcinoma cell lines [5,22,23]. In fact, in a rat pituitary somatotroph cell line, ghrelin exerts a proliferative effect via the mitogen-activated protein kinase (MAPK) pathway [24]. The pre-antral to antral follicle transition marks an important developmental step during follicular development where a subset of the pre-antral granulosa cells, probably those directly associated with the oocyte, differentiate into cumulus cells with competence to undergo cumulus expansion after the preovulatory LH surge [25]. GnRH and LH are important regulators for folliculogenesis, luteinization, and establishment of pregnancy as well [26,27]. Hence, the roles of ghrelin in follicular development and survival may depend on gonadotropin status and developmental state of the follicle.
During the ovulatory cycle, the OSE is subject to a series of injury and repair processes associated with follicular rupture and CL formation, which involve natural inflammatory events [28]. Expression of ghrelin within the OSE, which we have confirmed here, supports the theory that ghrelin play a vital role in OSE cells [12]. OSE is an obligate component of the ovulatory process as it plays an essential role during follicle rupture at the ovarian surface and formation of CL. Following ovulation, the small luteal cells of the CL are thought to originate from the follicular theca cells, whereas the large luteal cells are of GC origin. To ascertain the physiological regulation of ghrelin gene expression in sheep ovarian CL, relative mRNA levels were assessed during the different stages of the estrous cycle and pregnancy. Interestingly, ghrelin mRNA levels varied significantly depending on the phase of the estrous cycle. Ghrelin mRNA levels were highest at the beginning of the formation and mature phases, and lower during regression. Notably, a dynamic pattern of ovarian ghrelin mRNA expression was described in rat showing that ghrelin mRNA level varied depending on the phase of the cycle, with the lowest expression level in the proestrus and maximal value in the diestrus phase [10]. In addition, ovarian CL ghrelin gene expression was monitored throughout pregnancy. In our analysis, expression levels of ghrelin were higher from very early-pregnancy to mid-pregnancy, and significantly decreased later in gestation. Most studies on the
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biochemistry and structure of the CL suggest that during late pregnancy the CL of the rat is at the very early stages of structural regression, with no changes at the morphological level, but with changes at the molecular level (functional regression). Our results supported this report despite differences in animal species. In conclusion, we have demonstrated that the peptide ghrelin is expressed in sheep ovarian follicles at all developmental stages as well as in functional CL. Dynamic changes in the profile of ovarian expression of ghrelin mRNA during the estrous cycle and pregnancy are highly suggestive of a finely tuned regulatory network. Overall, the presence of the ghrelin signaling system within the sheep ovary especially in the oocytes opens up the possibility of a potential regulatory role of this novel molecule in reproductive function. Acknowledgements This work was supported by grants from National Natural Science Foundation of China (30660128 and 30860201) and Natural Science Foundation of Inner Mongolia (200711020506). The authors thank Mr. Yongmiao Feng and Dr. Bo Tang for their assistance and technical support during the studies. We also thank Dr. Hua Shao for his help with revision of the manuscript. References [1] Kojima M, Hosoda H, Date Y, Nakazato M, Matsuo H, Kangawa K. Ghrelin is a growth-hormone-releasing acylated peptide from stomach. Nature 1999;402:656–60. [2] van der Lely AJ, Tschop M, Heiman ML, Ghigo E. Biological, physiological, pathophysiological, and pharmacological aspects of ghrelin. Endocr Rev 2004;25:426–57. [3] Gualillo O, Lago F, Gomez-Reino J, Casanueva FF, Dieguez C. Ghrelin, a widespread hormone: insights into molecular and cellular regulation of its expression and mechanism of action. FEBS Lett 2003;552:105–9. [4] Duxbury MS, Waseem T, Ito H, Robinson MK, Zinner MJ, Ashley SW, et al. Ghrelin promotes pancreatic adenocarcinoma cellular proliferation and invasiveness. Biochem Biophys Res Commun 2003;309:464–8. [5] Xia Q, Pang W, Pan H, Zheng Y, Kang JS, Zhu SG. Effects of ghrelin on the proliferation and secretion of splenic T lymphocytes in mice. Regul Pept 2004;122:173–8. [6] Maccarinelli G, Sibilia V, Torsello A, Raimondo F, Pitto M, Giustina A, et al. Ghrelin regulates proliferation and differentiation of osteoblastic cells. J Endocrinol 2005;184:249–56. [7] Garcia MC, Lopez M, Alvarez CV, Casanueva F, Tena-Sempere M, Dieguez C. Role of ghrelin in reproduction. Reproduction 2007;133:531–40. [8] Harrison JL, Adam CL, Brown YA, Wallace JM, Aitken RP, Lea RG, et al. An immunohistochemical study of the localization and developmental expression of ghrelin and its functional receptor in the ovine placenta. Reprod Biol Endocrinol 2007;5:25.
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