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Testis |
Department of Cell Biology, Physiology, and Immunology,3 University of Córdoba, 14004 Córdoba, Spain
Departments of Physiology and Pediatrics,4 University of Turku, 20520 Turku, Finland
Centre for Molecular Biotechnology,5 Queensland University of Technology, Brisbane, Queensland, Australia
Departments of Medicine6
Physiology,7 University of Santiago de Compostela, 15705 Santiago de Compostela, Spain
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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follicle-stimulating hormone, ghrelin, growth hormone secretagogue, luteinizing hormone, receptor, testis
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Although most of the biological actions of ghrelin known to date are carried out at central levels, emerging evidence indicates that additional, as yet poorly characterized, peripheral actions of ghrelin are likely to take place. In this sense, a widespread pattern of expression of the genes encoding ghrelin and its cognate receptor has been reported very recently in humans [11], and GHS/ghrelin binding sites have been demonstrated in a variety of peripheral human tissues [12]. In addition, a number of noncentral tissues, such as placenta and kidney, have been shown to express ghrelin protein [13, 14]. However, the physiological relevance of ghrelin signaling in such peripheral systems remains to be fully established.
Recent data from our laboratory indicate that ghrelin and its cognate GHS-R are expressed in rat testis [15, 16]. Complete testicular function is critically dependent on the interaction of a plethora of endocrine, paracrine, and autocrine regulatory signals [17, 18], and a number of factors with pivotal roles in the regulation of the growth axis (e.g., growth hormone-releasing hormone and insulin-like growth factor-I) and body weight homeostasis (e.g., leptin) have been involved in the direct control of testis function [19–22]. Indeed, our initial evidence suggests that ghrelin may also participate in such a regulatory network [15]. However, the mode of action of ghrelin upon the male gonad as well as the potential regulatory mechanisms for the testicular effects of this molecule remain to be characterized.
In the present study, analysis of the pattern of expression of the GHS-R gene in rat testis was undertaken in different developmental and experimental settings in order to provide further insight into the testicular mode of action of ghrelin. Thus comparative evaluation of expression of total GHS-R and specific GHS-R1a mRNAs was conducted throughout postnatal development and at different stages of the spermatogenic cycle. To note, overall expression of the GHS-R gene (i.e., net levels of the mRNAs encoding GHS-R 1a and 1b forms) was defined as total GHS-R expression, whereas that of the messenger encoding the full-length 1a type was termed isoform-specific GHS-R1a expression. In addition, the pattern of cellular distribution of the GHS-R signal within adult testis tissue was assessed, and regulation of expression of GHS-R transcripts by homologous (i.e., ghrelin) and heterologous (i.e., gonadotropins) signals was analyzed. Overall, evidence for testicular expression of the GHS-R gene in a developmental, stage-specific, and hormonally regulated manner further supports the contention of a finely tuned, direct action of ghrelin in the functional control of rat testis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Wistar male rats bred in the vivarium of the University of Córdoba were used, unless otherwise stated. The day the litters were born was considered Day 1 of age. The animals were maintained under constant conditions of light (14L:10D; lights on 0700 h) and temperature (22°C) and were weaned at Day 21 of age in groups of five rats with free access to pelleted food and tap water. Experimental procedures were approved by the Córdoba University Ethical Committee for animal experimentation and were conducted in accordance with the European Union norm for care and use of experimental animals. Synthetic rat ghrelin, with n-octanoyl modification at Ser3, was obtained from Bachem AG (Bubendorf, Switzerland), and highly purified hCG (Profasi) was purchased from Serono (Madrid, Spain). Human recombinant FSH was kindly provided by Prof. J.A.F. Tresguerres (Gonal-F, Serono).
Experimental Designs
Assessment of expression of total GHS-R and specific GHS-R1a mRNAs in rat testis was carried out in different developmental and experimental settings. In experiment 1, comparative analysis of expression of the above targets was conducted at different stages of postnatal development, from neonatal period to adulthood. To this end, groups of animals (n = 5–10) were sequentially killed and testicular samples were obtained from 1-, 5-, 10-, 15-, 20-, 30-, 45-, and 75-day-old rats. Specific age-points were selected on the basis of previous references [23, and references therein], to correspond to neonatal (1-d), infantile (5-, 10-d), prepubertal (15-, 20-d), pubertal-early adult (30-, 45-d), and adult (75-d) stages of postnatal maturation. In addition, testicular tissue samples were taken from adult, 75-day-old animals, and processed for in situ hybridization and immunohistochemistry, as described below.
In experiment 2, expression of total GHS-R and specific GHS-R1a mRNAs was assessed in seminiferous tubule preparations at different stages of the spermatogenic cycle. Microdissection of seminiferous tubule segments of testes from Sprague-Dawley rats was carried out as described in detail elsewhere [24]. Briefly, testes from 75-day-old rats were decapsulated and 5 mm seminiferous tubule segments were isolated under a transilluminating stereomicroscope. Specific stages of the seminiferous epithelial cycle were identified as described previously [25] and pooled in four major groups corresponding to stages II–VI, stages VII–VIII, stages IX–XII, and stages XIII–I of the spermatogenic cycle. Microdissection of tubule segments was performed in DMEM-Ham's F-12 medium (1:1; Life Technologies, Paisley, U.K.) supplemented with 1.25 g/L sodium bicarbonate, 10 mg/L gentamicin sulfate, and 1 g/L BSA. After exhaustive washing, tubular tissue was processed for RNA analysis as described below.
Finally, a set of experiments was conducted to evaluate the potential regulation of testicular expression of GHS-R gene expression by homologous (i.e., the cognate ligand ghrelin) and heterologous (i.e., gonadotropins) signals. To this end, in experiment 3 expression of the total GHS-R and specific GHS-R1a mRNAs in rat testis tissue was evaluated after challenge with different doses of ghrelin in vitro. As experimental setting, slices of testicular tissue were obtained from 75-day-old rats and incubated for 180 min in the presence of increasing concentrations (10-9–10-7 M) of rat ghrelin, as described in detail elsewhere [15]. A similar experimental procedure has been previously used by our group to analyze the testicular effects of the adipocyte-derived hormone leptin in terms of testosterone secretion, steroidogenic gene expression, and leptin receptor regulation [21, 26, 27]. At the end of the 180-min incubation period, testicular samples were processed for RNA analysis as described below. In addition, in experiment 4 the regulation of testicular expression of the GHS-R gene by gonadotropins was explored in vivo. Thus expression levels of total GHS-R and specific GHS-R1a mRNAs were assayed in testes from adult intact rats injected at 1000 h with hCG (25 IU/rat) or human recombinant FSH (12.5 IU/rat) and sampled 2, 4, 8, and 24 h after administration. Paired vehicle-injected animals served as controls.
RNA Analysis by Semiquantitative RT-PCR
Total RNA was isolated from testis samples from the different experimental settings using the single-step, acid guanidinium thiocyanate-phenol-chloroform extraction method [28]. In the experimental groups, testicular expression of total GHS-R and isoform-specific GHS-R1a mRNAs was assessed by RT-PCR, optimized for semiquantitative detection. Such a method is especially useful for highly sensitive detection and discrimination between alternatively spliced species (e.g., isoform-specific GHS-R1a vs. total GHS-R transcripts). Evaluation of total GHS-R mRNA expression was carried out using a specific primer pair (forward [5'-AGG CAA CCT GCT CAC TAT GCT G-3'] and reverse [5'-GAC AAG GAT GAC CAG CTT CAC G-3']) flanking a 321-bp coding area of GHS-R cDNA common for both type 1a and 1b GHS-R forms. Expression levels of the mRNA specifically encoding type 1a GHS-R (i.e., the biologically active form of the receptor) were assessed using a type 1a-specific primer pair (forward [5'-TTC TTT CTA CCG GTC TTC TGC CTC-3'] and reverse [5'-GGA CAC CAG GTT GCA GTA CTG GCT-3']) spanning over the single intron of the GHS-R gene and allowing amplification of a 261-bp fragment of GHS-R cDNA unique to the type 1a form. The sets of primers were synthesized according to the published rat cDNA sequence of GHS-R [9] and selected on the basis of previous references [15, 29–31]. To note, amplification of total GHS-R species using the indicated primer pair inconsistently resulted in faint cogeneration of an amplicon of 
250-base pair (bp) length, together with a 321-bp band of expected size, in keeping with previous references [29, 30]. This transcript has been reported as related to GHS-R and has been stated as not to interfere with semiquantitative determination [29]; thus it was not considered for analysis. In addition, as internal control, amplification of a 290-bp fragment of L19 ribosomal protein mRNA was carried out in parallel in each sample using the following primer pair: L19 sense (5'-GAA ATC GCC AAT GCC AAC TC-3') and L19 as (5'-ACC TTC AGG TAC AGG CTG TG-3'), as described elsewhere [15, 21].
For amplification of the targets, RT and PCR were run in two separate steps. In addition, to enable appropriate amplification in the exponential phase for each target, PCR amplification of specific signals and L19 ribosomal protein transcripts was carried out in separate reactions with a different number of cycles, but using similar amounts of the corresponding cDNA templates, generated in single RT reactions, as previously described [15, 16, 21]. PCR reactions consisted in a first denaturing cycle at 97°C for 5 min, followed by a variable number of cycles of amplification defined by denaturation at 96°C for 1.5 min, annealing for 1.5 min, and extension at 72°C for 3 min. A final extension cycle of 72°C for 15 min was included. Annealing temperature was adjusted for each target: 60°C for total GHS-R, 62°C for GHS-R1a subtype, and 56°C for L19 ribosomal protein transcript. Different numbers of cycles were tested to optimize amplification in the exponential phase of PCR. On this basis, 32 and 23 PCR cycles were chosen for further analysis of GHS-R transcripts and L19 species, respectively (see Results).
PCR-generated DNA fragments were resolved in Tris-borate buffered 1.5% agarose gels and visualized by ethidium bromide staining. Specificity of PCR products was confirmed by direct sequencing (NewBiotechnic Ltd., Sevilla, Spain). In all assays, liquid controls and reactions without RT were included that yielded negative amplification (data not shown), thus ruling out the possibility of spurious amplification of the signals. When relevant, quantitative evaluation of RT-PCR signals was carried out by densitometric scanning using an image analysis system (1-D Manager; TDI Ltd., Madrid, Spain). The values for the specific targets were normalized to those of internal controls to express arbitrary units of relative abundance of the transcripts.
In Situ Hybridization
Five micrometer sections of adult (75-day-old) testis tissue were used for in situ hybridization. To assess the cellular distribution of GHS-R gene expression, a specific antisense 35S-labeled RNA probe complementary to an area of GHS-R cDNA unique to the type 1a form was synthesized using the Riboprobe system II kit (Promega, Madison, WI), T7 RNA polymerase, and [35S]-CTP. SalI-linearized template, constructed by subcloning a 261-bp fragment of GHS-R cDNA into a pGEM-T vector, was used. This cDNA was generated by RT-PCR using primer pair and conditions for isoform-specific GHS-R1a amplification, as described above. As control, adjacent sections were hybridized using a sense radio-labeled, cRNA probe generated as described above except for the use of a NcoI-linearized template and Sp6 RNA polymerase. Pretreatment and hybridization of sections were performed as described previously [24]. Finally, the slides were processed for liquid emulsion autoradiography using NTB-3 emulsion solution (Eastman Kodak). The sections were exposed at 4°C for 2–3 wk and developed at 12°C by treatment with D-19 solution (Eastman Kodak, Rochester, NY).
GHS-R1a Immunohistochemistry
Immunohistochemical detection of GHS-R1a peptide was carried out in 4% paraformaldehyde fixed sections of rat testes from adult (75-day-old) rats using a rabbit polyclonal antibody generated against a synthetic peptide corresponding to the C-terminal fragment (RAWTESSINTC) of the human GHS-R1a protein conjugated to diphtheria toxin (Mimotopes, Melbourne, Australia), as described in detail previously [32]. To note, Western analyses using this antibody identified a single specific band of approximately the predicted size (45 kDa) for the GHS-R1a in the ALVA-41 and DU145 prostate cancer cell lines (L.K.C., personal observation), which express the GHS-R1a mRNA isoform and GHS-R1a protein [32]. For immunolabeling, testicular sections (5 µm thick) were submitted to antigen retrieval in a microwave oven (2 x 5 min at 700 W) and incubated overnight with the primary anti-GHS-R1a antibody (diluted 1:200). The sections were then processed according to the avidin-biotin-peroxidase complex (ABC) technique, as described elsewhere [16]. Negative controls were run routinely in parallel by replacing the primary antibody by preimmune serum or PBS. In addition, as control for the specificity of GHS-R1a antibody, immunohistochemical reactions were carried out in testicular tissue following preabsorption of the antiserum overnight at 4°C with 1 mg/ml of the synthetic peptide (RAWTESSINTC), to which it was raised against. This procedure completely abolished immunolabeling of adult testis sections (see Fig. 4A).
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Presentation of Data and Statistics
RT-PCR analyses were carried out, at least in triplicate, using independent RNA samples. When relevant, semiquantitative RNA data are presented as mean ± SEM. Quantitative results were analyzed for statistically significant differences using ANOVA, followed by Tukey's test (P 
0.05 was considered significant).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To obtain optimal conditions for amplification (i.e., in the exponential phase of PCR), different numbers of cycles were tested for each target. As shown in Figure 1, plotting of intensity of PCR signals (as expressed by absolute OD values) against the number of amplification cycles revealed a strong linear relationship between cycles 24–39 in the case of GHS-R (correlation coefficient r2 = 0.985), GHS-R1a (coefficient r2 = 0.978), and cycles 17–29 in the case of L19 (coefficient r2 = 0.992). Thus PCR amplification of GHS-R-related and L19 ribosomal protein transcripts was conducted in separated reactions using 32 and 23 amplification cycles, respectively. The validity of the above RT-PCR assays for semiquantitative analysis is supported by 1) the selection, for each target, of amplification conditions in the exponential phase of PCR; 2) the repetitive observation of results within experimental groups (at least three assays per data-point using independent RNA and tissue samples); and 3) the use of an appropriate internal control.
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Testicular Expression of Total GHS-R and Isoform-Specific GHS-R1a mRNAs Along Postnatal Development
Assessment of expression levels of total GHS-R and isoform-specific GHS-R mRNAs was conducted in testicular samples throughout postnatal development, from the neonatal period to adulthood. Our semiquantitative analysis revealed persistent expression of total GHS-R mRNA in rat testis at all age points studied (1-, 5-, 10-, 15-, 20-, 30-, 45-, and 75-day-old rats) at rather constant relative levels, with maximum expression in adult (75-day-old) samples and minimum values in pubertal (45-day-old) testis tissue. In clear contrast, isoform-specific GHS-R1a mRNA expression remained undetectable in rat testis before Day 30 of age and sharply increased thereafter, with maximum expression values detected in adult 75-day-old testicular samples (Fig. 2).
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Pattern of Cellular Distribution of GHS-R Signal in Adult Testis Tissue
The pattern of cellular expression of the GHS-R gene within adult (75-day-old) testis tissue was evaluated by means of in situ hybridization using a specific antisense riboprobe recognizing the 1a form of GHS-R. Localization analyses revealed a scattered pattern of GHS-R1a mRNA expression, with deposition of hybridization signals in the seminiferous tubules and the interstitial space (Fig. 3). At higher magnification, specific signals were demonstrated in apparent Leydig and Sertoli cells, with possible location in germ cells also. In addition, assessment of the cellular distribution of GHS-R peptide in adult rat testis was conducted by immunohistochemistry using a specific polyclonal antibody raised against the C-terminal fragment of the type 1a GHS-R [32]. Such an analysis demonstrated the presence of GHS-R1a peptide in rat testis tissue. Moreover, in keeping with our in situ hybridization data, GHS-R1a immunoreactivity was mapped to interstitial Leydig cells, as well as to Sertoli cells within the seminiferous tubules (Fig. 4). Specificity of GHS-R1a immunolabeling was confirmed by preabsorption of the primary antibody with the synthetic peptide to which it was raised against; a procedure that completely blocked GHS-R1a immunoreactivity in adult testis sections.
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Stage-Specific Expression of Total GHS-R and Isoform-Specific GHS-R1a mRNAs in Seminiferous Tubule Preparations
Analysis of expression levels of total GHS-R and isoform-specific GHS-R1a mRNAs was undertaken in preparations of seminiferous tubule fragments isolated at different stages of the spermatogenic cycle. Upon microdissection of seminiferous tubule segments under a transilluminating stereomicroscope, specific stages of the seminiferous epithelial cycle were identified as described previously [25] and pooled in four major groups corresponding to stages II–VI, stages VII–VIII, stages IX–XII, and stages XIII–I of the spermatogenic cycle. In keeping with our in vivo data, positive amplification of total GHS-R and specific GHS-R1a mRNAs was obtained in tubule preparations at all stages of the spermatogenic cycle. However, the relative levels of expression of these signals disparately varied throughout the cycle. For the GHS-R transcript, minimum values were detected at stages VII–VIII, whereas maximum expression levels were observed at stages IX–XII. Such a peak in expression was not detected, however, for GHS-R1a mRNA that remained at lower expression levels at stages IX–XII than at stages II–VI and XIII–I (Fig. 5). For both signals, moderate to high expression levels were detected at stages XIII–VI of the spermatogenic cycle.
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Homologous and Heterologous Regulation of Total GHS-R and Isoform-Specific GHS-R1a mRNA Expression in Rat Testis
In a next step, the ability of different hormonal signals to regulate testicular expression of total GHS-R and isoform-specific GHS-R1a mRNAs was evaluated. To this end, in a first approach, analysis of homologous regulation (i.e., by the ligand of the cognate receptor) was undertaken in adult rats using an in vitro setting. RT-PCR assays confirmed high expression levels of total GHS-R and specific GHS-R1a transcripts in testicular samples dissected out from adult rats immediately after decapitation. These values were taken as reference basal expression levels. Short-term (4-h) incubation of testis tissue in serum-free (SF) medium resulted in a significant decrease in the relative mRNA levels of both GHS-R and GHS-R1a transcripts. Such responses were completely prevented by 3-h incubation in the presence of increasing concentrations of ghrelin (10-9–10-7 M), but not of hCG (10 IU/ml), thus suggesting specificity for the up-regulatory effect of ghrelin (Fig. 6). Interestingly, subtle differences were noted in the dose-dependency for the above stimulatory responses, as total GHS-R mRNA levels were similarly elevated over basal SF-medium values by all doses of ghrelin (10-9–10-7 M) tested, whereas expression of isoform-specific GHS-R1a species was selectively increased by challenge with 10-8–10-7 M ghrelin, but not 10-9 M ghrelin (Fig. 6).
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In addition, the effects of exposure to hCG (as super-agonist of LH) and recombinant FSH upon testicular expression of total GHS-R and isoform-specific GHS-R1a mRNAs were explored in vivo. Assessment of expression levels of both signals was conducted 2, 4, 8, and 24 h after administration of gonadotropins. This analysis revealed that exposure to 12.5 IU of recombinant FSH induced an increase in testicular expression levels of total GHS-R mRNA, which was statistically significant at 8 and 24 h following FSH injection. In contrast, administration of 25 IU of highly purified hCG did not alter the relative expression levels of total GHS-R mRNA in testis tissue at any time point studied (Fig. 7). Similarly, recombinant FSH treatment significantly stimulated isoform-specific GHS-R1a mRNA expression in rat testis along the study period (Fig. 8), whereas hCG was ineffective (data not shown). However, the time course of response to recombinant FSH in terms of total GHS-R and specific GHS-R1a expression was partially different, as significant stimulation of GHS-R1a mRNA expression was observed at all time points (2–24 h) studied.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Our semiquantitative analysis demonstrated persistent expression of total GHS-R mRNA in rat testis at rather constant relative levels throughout different stages of postnatal development, including the neonatal, infantile, prepubertal, pubertal-early adult, and adult periods. In clear contrast, isoform-specific GHS-R1a mRNA expression was undetectable during the neonatal, infantile, and prepubertal stages of postnatal development and sharply increased from Day 30 onward, reaching maximum expression values in the adult age. Thus comparative analysis of the profile of expression of both signals suggests that during pubertal development a shift in the pattern of splicing of the GHS-R gene takes place in rat testis that favors expression of the biologically active type 1a form of the receptor. Likely, in earlier stages of postnatal development the predominant receptor form is the truncated GHS-R1b type. The relevance of such a phenomenon is presently under investigation, but considering that the functional capacities of both receptor forms are strikingly different [7], it is reasonable to propose that net ghrelin sensitivity changes significantly along postnatal testicular development.
Expression of the GHS-R gene in rat testis was further confirmed by in situ hybridization data, showing a scattered distribution of GHS-R1a mRNA within adult testis tissue that apparently included Sertoli cells within the seminiferous tubules and interstitial Leydig cells. This expression pattern was roughly similar to that observed at the peptide level by means of immunohistochemistry using a specific polyclonal antibody raised against the C-terminal fragment of the type 1a GHS-R. The presence of the functional GHS-R type 1a in Leydig cells is in keeping with the direct effect of ghrelin upon stimulated testosterone secretion in vitro [15]. However, our localization analyses clearly demonstrate that interstitial Leydig cells are not the only source of testicular GHS-R. Indeed, these data are in good agreement with our previous results on the persistent expression of GHS-R mRNA in rat testis after selective elimination of mature Leydig cells by administration of the toxicant ethylene dimethane sulfonate (EDS) [15]. Accordingly, expression of the GHS-R signal within the seminiferous tubules was detected (see Figs. 3 and 4). In addition, positive amplification of total GHS-R and specific GHS-R1a mRNAs was obtained in tubule preparations at all stages of the spermatogenic cycle, thus confirming expression of the GHS-R gene in the seminiferous epithelium. Interestingly, however, the relative levels of expression of total GHS-R and specific GHS-R1a signals disparately varied throughout the cycle. In detail, moderate to high expression levels were observed for both transcripts at stages XIII–I and II–VI, whereas minimum relative levels were detected at stages VII–VIII. To note, stages VII–VIII are highly androgen-dependent but are defined by a low sensitivity to FSH [36]. This may be related to the observed decrease in relative expression levels of total GSH-R and specific GHS-R1a mRNAs at these stages, as these transcripts were strongly up-regulated by FSH in vivo. In addition, a divergent pattern of expression between total GHS-R and isoform-specific GHS-R1a became apparent at stages IX–XI. At these stages, the expression levels of total GHS-R mRNA reached maximum values while specific GHS-R1a transcript remained at lower levels than those of stages XIII–VI. The relevance of such a divergent profile of expression in the cyclic function of Sertoli cells remains to be evaluated. Taken together, our current results demonstrate that the GHS-R gene is expressed in rat seminiferous epithelium in a stage-specific manner and suggest that changes in the pattern of splicing of the gene take place at certain phases of the spermatogenic cycle.
Regulation of GHS-R expression is likely a key event in the modulation of the biological actions of ghrelin in target tissues. Indeed, compelling evidence indicates that a plethora of heterologous endocrine signals modulate pituitary and hypothalamic expression of the GHS-R gene. Thus, total GHS-R mRNA levels are inhibited by GH [29, 33], whereas its expression is up-regulated by GHRH [34]. Additional regulatory factors are sex steroids and glucocorticoids [30, 35]. Similarly, homologous regulation is likely to take place as evidenced by down-regulation of total GHS-R mRNA expression in rat pituitary after exposure to the GH secretagogue L629,585 in vivo [34]. In our experiments, we assessed the hormonal regulation of GHS-R expression in rat testis. To this end, the ability of ghrelin and gonadotropins to modulate testicular expression of total GHS-R and isoform-specific GHS-R1a mRNAs was explored using in vitro and in vivo systems. Regulation of GHS-R mRNA expression by gonadotropins was evaluated given the pivotal role of these signals in the hormonal control of testicular function [37]. Our present results indicate that, as is the case at the pituitary and hypothalamus, testicular expression of the GHS-R gene is also under the regulation of homologous and heterologous signals, as ghrelin and FSH were able to coordinately increase both total GHS-R and isoform-specific GHS-R1a mRNA expression in rat testis.
Concerning homologous regulation, our analysis, using a static incubation setting, demonstrated that exposure to increasing doses (10-9–10-7 M) of the cognate ligand, ghrelin, significantly enhanced expression of total GHS-R and specific GHS-R1a transcripts over basal values, whereas short-term incubation in serum-free (SF) medium caused a significant decrease in the level of expression of both signals. It is noteworthy that the range of doses of ghrelin used in our in vitro setting is close to that of plasma ghrelin levels in basal and stimulated conditions in the rat [38, 39]. These observations are in agreement with the hypothesis that threshold ghrelin levels are required to maintain elevated expression of the GHS-R gene in adult rat testis. Accordingly, high mRNA levels of total GHS-R and specific GHS-R1a are detected in adult rat testis under in vivo conditions, with high circulating ghrelin levels, and after challenge with increasing concentrations of ghrelin in vitro, but low expression of the transcripts is observed after SF medium incubation. Interestingly, ligand-dependent modulation of GHS-R transcript in another steroidogenic tissue, such as the adrenal gland, takes place in a strikingly opposite manner, whereas a pattern of homologous regulation of GHS-R mRNA expression roughly analogous to that of the testis has been observed by our group in the pituitary [40]. Thus, regulation of GHS-R mRNA expression by homologous signals appears to be a widespread phenomenon. However, the precise nature of such an event is likely tissue-specific, which might reflect the divergent physiological roles of GHS-R signaling in different target systems.
In addition, our data indicate that, besides its cognate ligand, testicular expression of total GHS-R and isoform-specific GHS-R1a mRNAs is under the control of pituitary FSH. Considering that FSH receptors are solely expressed in Sertoli cells within the tubular compartment of the testis [37], it is possible that the stimulatory effect of recombinant FSH upon GHS-R transcripts may reflect a genuine direct action of this gonadotropin upon Sertoli cells. Alternatively, an indirect action of FSH upon other cell types within the seminiferous epithelium cannot be ruled out. Notably, direct stimulation of Leydig cells by administration of hCG (as super-agonist of LH) failed to alter both total GHS-R and specific GHS-R1a mRNA expression in rat testis, despite the fact that LH receptors are solely expressed in interstitial Leydig cells [37], and this cell type does express the GHS-R gene. However, secondary effects are possible, as LH/CG is the major stimulator of testosterone secretion by Leydig cells, and Sertoli cells are targets of testosterone action [37]. Indeed, a transient decline in testicular expression levels of GHS-R1a mRNA was associated with a decrease in serum testosterone levels following EDS treatment (unpublished data). The fact, however, that GHS-R transcripts remained unaltered after hCG stimulation in vivo strongly suggests that testosterone may play a permissive role, if any, in the control of GHS-R expression in rat testis. In addition, hormonal regulation of testicular GHS-R mRNA expression may involve interactions between different modulatory signals (e.g., ghrelin and FSH), as well as additional hormonal factors. From a general standpoint, evidence for the stage-specific expression of the mRNA encoding the functional 1a form of GHS-R in rat seminiferous tubules, under the regulation of pituitary FSH, strongly suggests that ghrelin signaling might be involved in the functional regulation of the tubular compartment of the testis. Further experimental work is in progress in our laboratory to elucidate the mode of action and functional role of ghrelin in rat seminiferous epithelium.
In the last few years, ghrelin has been identified as a novel neuroendocrine signal involved in the physiological control of GH secretion and food intake [1–6]. More recently, an "unexpected" reproductive facet of ghrelin has emerged. In this context, our group has shown recently that ghrelin is expressed in rat testis under the control of pituitary LH, and it is able to inhibit stimulated testosterone secretion in vitro [15, 16]. Moreover, a specific ghrelin gene-derived transcript has been identified in mouse testis [41]. In the present paper, comparative analysis of the expression profile of total GHS-R and isoform-specific GHS-R1a mRNAs in rat testis was undertaken in a number of experimental settings. All together, our data are the first to demonstrate that testicular expression of GHS-R mRNAs takes place in a developmental, stage-specific, and hormonally regulated manner; a phenomenon highly suggestive of a finely regulated, direct action of ghrelin in the functional control of rat testis. Disparate expression patterns of total GHS-R and type 1a specific mRNAs were observed at certain stages of postnatal development and spermatogenic cycle, thus raising the possibility that, in addition to net changes in GHS-R gene expression, the balance between receptor subtypes may represent a novel mechanism for the tuning of ghrelin sensitivity in rat testis.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Manuel Tena-Sempere, Physiology Section, Department of Cell Biology, Physiology and Immunology, Faculty of Medicine, University of Córdoba, Avda. Menéndez Pidal s/n, 14004 Córdoba, Spain. FAX: 34 957 218288; fi1tesem{at}uco.es
Received: 27 June 2002.
First decision: 17 July 2002.
Accepted: 22 November 2002.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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M. C Garcia, M. Lopez, C. V Alvarez, F. Casanueva, M. Tena-Sempere, and C. Dieguez Role of ghrelin in reproduction Reproduction, March 1, 2007; 133(3): 531 - 540. [Abstract] [Full Text] [PDF]
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 T. Ishikawa, H. Fujioka, T. Ishimura, A. Takenaka, and M. Fujisawa Ghrelin Expression in Human Testis and Serum Testosterone Level J Androl, March 1, 2007; 28(2): 320 - 324. [Abstract] [Full Text] [PDF]
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 C.-M. Yeung, C.-B. Chan, N. Y S Woo, and C. H K Cheng Seabream ghrelin: cDNA cloning, genomic organization and promoter studies. J. Endocrinol., May 1, 2006; 189(2): 365 - 379. [Abstract] [Full Text] [PDF]
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 K. J. Pulman, W. M. Fry, G. T. Cottrell, and A. V. Ferguson The Subfornical Organ: A Central Target for Circulating Feeding Signals J. Neurosci., February 15, 2006; 26(7): 2022 - 2030. [Abstract] [Full Text] [PDF]
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F. Gaytan, C. Morales, M. L. Barreiro, P. Jeffery, L. K. Chopin, A. C. Herington, F. F. Casanueva, E. Aguilar, C. Dieguez, and M. Tena-Sempere 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., March 1, 2005; 90(3): 1798 - 1804. [Abstract] [Full Text] [PDF]
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 P. L. Jeffery, R. P. Duncan, A. H. Yeh, R. A. Jaskolski, D. S. Hammond, A. C. Herington, and L. K. Chopin Expression of the Ghrelin Axis in the Mouse: An Exon 4-Deleted Mouse Proghrelin Variant Encodes a Novel C Terminal Peptide Endocrinology, January 1, 2005; 146(1): 432 - 440. [Abstract] [Full Text] [PDF]
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T. R. Kumar Divide and Differentiate: Ghrelin Instructs the Leydig Cells Endocrinology, November 1, 2004; 145(11): 4822 - 4824. [Full Text] [PDF]
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M. L. Barreiro, F. Gaytan, J. M. Castellano, J. S. Suominen, J. Roa, M. Gaytan, E. Aguilar, C. Dieguez, J. Toppari, and M. Tena-Sempere Ghrelin Inhibits the Proliferative Activity of Immature Leydig Cells in Vivo and Regulates Stem Cell Factor Messenger Ribonucleic Acid Expression in Rat Testis Endocrinology, November 1, 2004; 145(11): 4825 - 4834. [Abstract] [Full Text] [PDF]
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M. L. Barreiro, R. Pineda, V. M. Navarro, M. Lopez, J. S. Suominen, L. Pinilla, R. Senaris, J. Toppari, E. Aguilar, C. Dieguez, et al. Orexin 1 Receptor Messenger Ribonucleic Acid Expression and Stimulation of Testosterone Secretion by Orexin-A in Rat Testis Endocrinology, May 1, 2004; 145(5): 2297 - 2306. [Abstract] [Full Text] [PDF]
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F. Gaytan, M. L. Barreiro, J. E. Caminos, L. K. Chopin, A. C. Herington, C. Morales, L. Pinilla, R. Paniagua, M. Nistal, F. F. Casanueva, et al. Expression of Ghrelin and Its Functional Receptor, the Type 1a Growth Hormone Secretagogue Receptor, in Normal Human Testis and Testicular Tumors J. Clin. Endocrinol. Metab., January 1, 2004; 89(1): 400 - 409. [Abstract] [Full Text] [PDF]
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K. G. Mountjoy, C.-S. Jenny Wu, L. M. Dumont, and J. M. Wild Melanocortin-4 Receptor Messenger Ribonucleic Acid Expression in Rat Cardiorespiratory, Musculoskeletal, and Integumentary Systems Endocrinology, December 1, 2003; 144(12): 5488 - 5496. [Abstract] [Full Text] [PDF]
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