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BOR - Papers in Press, published online ahead of print November 27, 2002.
Biol Reprod 2002, 10.1095/biolreprod.102.008862
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BIOLOGY OF REPRODUCTION 68, 1631–1640 (2003)
DOI: 10.1095/biolreprod.102.008862
© 2003 by the Society for the Study of Reproduction, Inc.

Testis

Developmental, Stage-Specific, and Hormonally Regulated Expression of Growth Hormone Secretagogue Receptor Messenger RNA in Rat Testis1

M.L. Barreiro3, J.S. Suominen4, F. Gaytán3, L. Pinilla3, L.K. Chopin5, F.F. Casanueva6, C. Diéguez7, E. Aguilar3, J. Toppari4, and M. Tena-Sempere2,3

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


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Recent evidence from our research suggested the direct role of ghrelin in the control of testicular function. However, the pattern of expression and hormonal regulation of the gene encoding its cognate receptor (i.e., the growth hormone-secretagogue receptor [GHS-R]) in the male gonad remains to be fully elucidated. In this paper, overall expression of GHS-R mRNA in rat testis was compared with that of the functional receptor form, namely GHS-R type 1a, in different developmental and experimental settings. In addition, cellular distribution of GHS-R within adult testis tissue was assessed. Our analyses demonstrated persistent expression of the GHS-R gene in rat testis throughout postnatal development. In contrast, testicular expression of GHS-R type 1a mRNA remained undetectable before puberty and sharply increased thereafter. In adult testis, GHS-R1a mRNA expression presented a scattered pattern of cellular distribution, including Sertoli and Leydig cells that also showed specific GHS-R1a immunoreactivity. Expression of total GHS-R and specific GHS-R1a mRNAs was detected in isolated seminiferous tubule preparations, with varying levels throughout the defined stages of the spermatogenic cycle. In addition, testicular expression of total GHS-R and GHS-R1a mRNAs was up-regulated by exposure to ghrelin in vitro and after stimulation with FSH in vivo. In conclusion, our data demonstrate that expression of the GHS-R gene in rat testis takes place in a developmental, stage-specific, and hormonally regulated manner. Divergent expression of total GHS-R and type 1a specific mRNAs was detected 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.

follicle-stimulating hormone, ghrelin, growth hormone secretagogue, luteinizing hormone, receptor, testis


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Ghrelin is a 28-amino acid peptide, with an essential n-octanoyl modification at Ser3, recently identified as the endogenous ligand for the growth hormone-secretagogue receptor (GHS-R) [1, 2]. This peptide is primarily expressed in the stomach and hypothalamus and it is likely involved in the physiological network controlling GH secretion and food intake [1–6]. Its cognate receptor, the GHS-R, belongs to the large family of G-protein coupled, seven transmembrane spanning receptors [7–9], it being mainly expressed at central neuroendocrine tissues as the pituitary and hypothalamus [10]. Two GHS-R subtypes, generated by alternative splicing of a single gene, have been identified: the full-length type 1a receptor and the truncated GHS-R type 1b [8, 9]. The GHS-R1a is the functionally active, signal-transducing form of the receptor. In contrast, the GHS-R1b lacks the transmembrane domains 6 and 7, and it is apparently devoid of high-affinity ligand binding and signal transduction capacity [7]. Whether this subtype plays a role in the functional regulation of GHS-R1a remains to be solved [7], but it is possible that overall ghrelin sensitivity in target tissues may depend, at least partially, on the balance of expression between GHS-R isoforms.

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.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals and Drugs

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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  		FIG. 4. Immunolocalization of GHS-R1a protein in adult rat (75-day-old) testis sections. A) A representative control reaction carried out after preabsorption of primary anti-GHS-R1a antibody with the synthetic peptide it was raised against. B) An adjacent section after immunolabeling with the specific primary antibody. Interstitial Leydig cells are indicated by filled arrows, whereas within seminiferous tubules Sertoli cells are indicated by open arrows. At higher magnification (C), specific GHS-R1a immuno-reactivity was clearly located upon Leydig and Sertoli cells. Scale bars in (A) and (B) = 50 µm; in (C) = 20 µm

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).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Optimization of Semiquantitative RT-PCR Assays

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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  		FIG. 1. Optimization of RT-PCR conditions for semiquantitative evaluation of relative expression levels of the target mRNA species. For amplification in the exponential phase of PCR, different numbers of cycles were tested for each transcript. Quantitative analysis of the cycle-dependency for the generated PCR signals demonstrated a strong linear relationship between cycles 24 to 39 in the case of total GHS-R (correlation coefficient r2 = 0.985) and specific GHS-R1a subtype (correlation coefficient r2 = 0.978), and cycles 18 to 27 in the case of L19 ribosomal protein (correlation coefficient r2 = 0.992). Optimization of RT-PCR conditions was carried out using total RNA from control 75-day-old rat testis. Values are given as mean ± SD of two independent determinations. A representative ethidium bromide-stained gel electrophoresis of the DNA products generated for each target is presented in the insets.

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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  		FIG. 2. Developmental profile of expression of total GHS-R and specific GHS-R1a mRNAs in rat testis throughout the stages of postnatal maturation. Representative RT-PCR assays of expression levels of GHS-R and GHS-R1a mRNAs in testicular samples from 1-, 5-, 10-, 15-, 20-, 30-, 45-, and 75-day-old rats are presented (upper panels). Parallel amplification of L19 ribosomal protein mRNA served as internal control. A 50-bp molecular weight marker (M) was used. Semiquantitative data on the expression levels of the targets in rat testis along postnatal development are shown (lower panels). Relative expression levels were obtained in each sample by normalization of absolute optical densities (ODs) of the specific target to that of the L19 signal. For presentation of data, maximum expression levels along the study period were taken as 100% and the others were normalized accordingly, thus allowing semiquantitative comparison. Values are the mean ± SEM of at least three independent determinations. Groups with different superscript letters are statistically different (P < 0.01; ANOVA followed by Tukey's test)

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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  		FIG. 3. Representative photomicrographs of GHS-R1a mRNA in situ hybridization in adult rat testis. Testicular sections from 75-day-old rats were hybridized with a specific antisense RNA probe recognizing the 1a form of GHS-R, as described in Materials and Methods (upper panels). Specific hybridization signals for GHS-R1a mRNA were localized in interstitial Leydig cells (filled arrows) and, within seminiferous tubules, to apparent Sertoli cells (open arrows), with possible specific deposition of silver grains in germ cells also. Adjacent sections hybridized with the corresponding control sense cRNA probe are also shown (lower panels). Brightfield (A and B) and darkfield (A' and B') photomicrographs are presented. The emulsion-coated slides were exposed in the dark for 2–3 wk at 4°C. Scale bars = 50 µm

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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  		FIG. 5. Expression profile of total GHS-R and specific GHS-R1a mRNAs in seminiferous tubule segments at different stages of the spermatogenic cycle. Upper panels: representative RT-PCR assays of expression levels of GHS-R and GHS-R1a mRNAs in tubule preparations isolated at stages II–VI, VII–VIII, IX–XII, and XIII–I of the spermatogenic cycle, as described in Materials and Methods. Parallel amplification of L19 ribosomal protein mRNA served as internal control. A 50-bp molecular weight marker (M) was used. Lower panels: semiquantitative data on the expression levels of the targets in the different stages. Relative levels were obtained in each sample by normalization of absolute ODs of the specific targets to that of the L19 signal. For presentation of data, expression levels in stages II–VI were arbitrarily taken as 100% and the others were normalized accordingly, thus allowing semiquantitative comparison. Values are the mean ± SEM of at least four independent determinations. Groups with different superscript letters are statistically different (P < 0.01; ANOVA followed by Tukey's test)

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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  		FIG. 6. Pattern of homologous regulation of total GHS-R and isoform-specific GHS-R1a mRNA expression in rat testis. Upper panels: representative RT-PCR assays of the expression levels of GHS-R and GHS-R1a transcripts in testicular samples in basal in vivo conditions (T), and after short-term (4-h) incubation either in serum-free medium (0) or medium containing increasing concentrations of ghrelin (10-9–10-7 M). In addition, expression levels of the targets in testis samples after challenge with hCG (10 IU/ml) in vitro are presented. Parallel amplification of L19 ribosomal protein mRNA served as internal control. A 50-bp molecular weight marker (M) was used. Lower panels: semiquantitative data are shown on the steady-state levels of GHS-R and GHS-R1a mRNAs in testicular samples from the above experimental groups. Relative expression levels were obtained, in each sample, by normalization of absolute ODs of the specific target to that of the L19 signal. For presentation of data, for each target, maximum expression values (T) were taken as 100% and the others were normalized accordingly, thus allowing semiquantitative comparison. Values are the mean ± SEM of at least three independent determinations. Groups with different superscript letters are statistically different (P < 0.01; ANOVA followed by Tukey's test)

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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  		FIG. 7. Pattern of regulation of total GHS-R mRNA expression in rat testis by heterologous signals (i.e., gonadotropins). Upper panels: representative RT-PCR assays of the testicular expression levels of GHS-R transcript in adult rats at different time points (2–24 h) after exposure to 25 IU hCG or 12.5 IU recombinant FSH in vivo. Parallel amplification of L19 ribosomal protein mRNA served as internal control. A 50-bp molecular weight marker (M) was used. Lower panels: semiquantitative data are shown on the steady-state levels of GHS-R mRNA in testicular samples from the above experimental groups. Relative expression levels were obtained, in each sample, by normalization of absolute ODs of the specific target to that of the L19 signal. For presentation of data, expression levels in control (0) samples (i.e., from vehicle-injected animals) were taken as 100% and the others were normalized accordingly, thus allowing semiquantitative comparison. Values are the mean ± SEM of at least three independent determinations. Groups with different superscript letters are statistically different (P < 0.01; ANOVA followed by Tukey's test)



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  		FIG. 8. Regulation of isoform-specific GHS-R1a mRNA expression in rat testis by FSH in vivo. Upper panels: a representative RT-PCR assay of the testicular expression levels of GHS-R1a transcript in adult rats at different time points (2–24 h) after exposure to 12.5 IU recombinant FSH in vivo. Parallel amplification of L19 ribosomal protein mRNA served as internal control. A 50-bp molecular weight marker (M) was used. Lower panel: semiquantitative data are shown on the steady-state levels of GHS-R1a mRNA in testicular samples from the above experimental groups. Relative expression levels were obtained, in each sample, by normalization of absolute ODs of the specific target to that of the L19 signal. For presentation of data, expression levels in control (0) samples (i.e., from vehicle-injected animals) were taken as 100% and the others were normalized accordingly, thus allowing semiquantitative comparison. Values are the mean ± SEM of at least three independent determinations. Groups with different superscript letters are statistically different (P < 0.01; ANOVA followed by Tukey's test)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The hallmark actions of the novel acylated peptide ghrelin (i.e., stimulation of GH secretion and food intake) are carried out at central neuroendocrine levels through interaction with its specific receptor. Notably, the mechanisms for modulation of ghrelin sensitivity in such target tissues remain to be fully elucidated. However, compelling evidence indicates that pituitary and hypothalamic expression of the GHS-R gene appears as a tightly regulated event, where both inhibitory and stimulatory signals interact to ensure adequate expression levels [29, 30, 33–35]. In addition, the balance between the functional type 1a and the truncated type 1b forms of the GHS-R might play a role in this phenomenon [7]. Present results confirm and extend our previous observations on the expression of the GHS-R gene in rat testis throughout postnatal development [15] and are the first to demonstrate expression of the biologically active GHS-R form (i.e., the type 1a receptor) in rat testicular tissue from puberty onward. Moreover, novel evidence is presented for the stage-specific and hormonally regulated expression of the GHS-R gene in rat testis. Overall, our current data strongly suggest that, as is the case at central neuroendocrine levels, testicular expression of GHS-R is a finely tuned event that may play a pivotal role in the modulation of ghrelin sensitivity in rat testis.

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.


   ACKNOWLEDGMENTS
 
The authors are indebted to Rocío Campón for her excellent technical assistance.


   FOOTNOTES
 
1 Supported by grant PM-98-0163 from DGESIC (Ministerio de Ciencia y Tecnología, Spain); project 1FD97-0696-02 (FEDER); and grants from the Academy of Finland and Turku University Central Hospital. Back

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 Back

Received: 27 June 2002.

First decision: 17 July 2002.

Accepted: 22 November 2002.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Kojima M, Hosoda H, Date Y, Nakazato M, Matsuo H, Kangawa K. Ghrelin is a growth-hormone acylated peptide from stomach. Nature 1999 402:656-666[CrossRef][Medline]
  2. Kojima M, Hosoda H, Matsuo H, Kangawa K. Ghrelin: discovery of the natural endogenous ligand for the growth hormone secretagogue receptor. Trends Endocrinol Metab 2001 12:118-122[CrossRef][Medline]
  3. Takaya K, Ariyasu H, Kanamoto N, Iwakura H, Yoshimoto A, Harada M, Mori K, Komatsu Y, Usui T, Shimatsu A, Ogawa Y, Hosoda K, Akamizu T, Kojima M, Kangawa K, Nakao K. Ghrelin strongly stimulates growth hormone release in humans. J Clin Endocrinol Metab 2000 85:4908-4911[Abstract/Free Full Text]
  4. Wren AM, Small CJ, Ward HL, Murphy KG, Dakin CL, Taheri S, Kennedy AR, Roberts GH, Morgan DG, Ghatei MA, Bloom SR. The novel hypothalamic peptide ghrelin stimulates food intake and growth hormone secretion. Endocrinology 2000 141:4325-4328[Abstract/Free Full Text]
  5. Tschop M, Smiley DL, Heiman ML. Ghrelin induces adiposity in rodents. Nature 2000 407:908-913[CrossRef][Medline]
  6. Arvat E, Maccario M, di Vito L, Broglio F, Benso A, Gottero C, Papotti M, Muccioli G, Dieguez C, Casanueva FF, Deghenghi R, Camanni F, Ghigo E. Endocrine activities of ghrelin, a natural growth hormone secretagogue (GHS), in humans: comparison and interactions with hexarelin, a nonnatural peptidyl GHS, and GH-releasing hormone. J Clin Endocrinol Metab 2001 86:1169-1174[Abstract/Free Full Text]
  7. Smith RG, van der Ploeg LHT, Howard AD, Feighner SD, Cheng K, Hickey GJ, Wyvratt MJ, Fisher MH, Nargund RP, Patchett AA. Peptidomimetic regulation of growth hormone secretion. Endocr Rev 1997 18:621-645[Abstract/Free Full Text]
  8. Howard AD, Feighner SC, Cully DF, Arena JP, Liberator PA, Rosenblum CI, Hamelin M, Hreniuk DL, Palyha OC, Anderson J, Paress PS, Diaz C, Chou M, Liu KK, McKee KK, Pong SS, Chaung LY, Elbrecht A, Dashkevicz M, Heavens R, Rigby M, Sirinathsinghji DJ, Dean DC, Melillo DG, Patchett AA, Nargund RP, Griffin PR, DeMartino JA, Gupta SK, Schaeffer JM, Smith RG, van der Ploeg LHT. A receptor in pituitary and hypothalamus that functions in growth hormone release. Science 1996 273:974-977[Abstract]
  9. McKee KK, Palyha OC, Feighner SD, Hreniuk DL, Tan CP, Phillips MS, Smith RG, van der Ploeg LH, Howard AD. Molecular analysis of rat pituitary and hypothalamic growth hormone secretagogue receptors. Mol Endocrinol 1997 11:415-423[Abstract/Free Full Text]
  10. Guan XH, Yu H, Palyha OC, McKee KK, Feighner SD, Sirinathsinghji DJ, Smith RG, van der Ploeg LH, Howard AD. Distribution of mRNA encoding the growth hormone secretagogue receptor in brain and peripheral tissues. Brain Res Mol Brain Res 1997 48:23-29[Medline]
  11. Gnanapavan S, Kola B, Bustin SA, Morris DG, McGee P, Fairclough P, Bhattacharya S, Carpenter R, Grossman AB, Korbonits M. The tissue distribution of the mRNA of ghrelin and subtypes of its receptor, GHS-R, in humans. J Clin Endocrinol Metab 2002 87:2988-2992[Abstract/Free Full Text]
  12. Papotti M, Ghé C, Cassoni P, Catapano F, Deghenghi R, Ghigo E, Muccioli G. Growth hormone secretagogue binding sites in peripheral human tissues. J Clin Endocrinol Metab 2000 85:3803-3807[Abstract/Free Full Text]
  13. Gualillo O, Caminos JE, Blanco M, García-Caballero T, Kojima M, Kangawa K, Dieguez C, Casanueva F. Ghrelin, a novel placental-derived hormone. Endocrinology 2001 142:788-794[Abstract/Free Full Text]
  14. Mori K, Yoshimoto A, Takaya K, Hosoda K, Ariyasu H, Yahata K, Mukoyama M, Sugawara A, Hosoda H, Kojima M, Kangawa K, Nakao K. Kidney produces a novel acylated peptide, ghrelin. FEBS Lett 2000 486:213-216[CrossRef][Medline]
  15. Tena-Sempere M, Barreiro ML, Gonzalez LC, Gaytan F, Zhang FP, Caminos JE, Pinilla L, Casanueva FF, Dieguez C, Aguilar E. Novel expression and functional role of ghrelin in rat testis. Endocrinology 2002 143:717-725[Abstract/Free Full Text]
  16. Barreiro ML, Gaytan F, Caminos JE, Pinilla L, Casanueva FF, Aguilar E, Dieguez C, Tena-Sempere M. Cellular location and hormonal regulation of ghrelin expression in rat testis. Biol Reprod 2002; 67:1768–1776.
  17. Saez JM. Leydig cell: endocrine, paracrine, and autocrine regulation. Endocr Rev 1994 15:574-626[CrossRef][Medline]
  18. Skinner MK. Cell-cell interactions in the testis. Endocr Rev 1991 12:45-77[Medline]
  19. Ciampani T, Fabbri A, Isidori A, Dufau ML. Growth hormone-releasing hormone is produced by rat Leydig cells in culture and acts as a positive regulator of Leydig cell function. Endocrinology 1992 131:2785-2792[Abstract]
  20. Baker J, Hardy MP, Zhou J, Bondy C, Lupu F, Bellve AR, Efstratiadis A. Effects of an Igf1 gene null mutation on mouse reproduction. Mol Endocrinol 1996 10:903-918[Abstract]
  21. Tena-Sempere M, Pinilla L, Zhang FP, Gonzalez LC, Huhtaniemi I, Casanueva FF, Dieguez C, Aguilar E. Developmental and hormonal regulation of leptin receptor (Ob-R) messenger ribonucleic acid expression in rat testis. Biol Reprod 2001 64:634-643[Abstract/Free Full Text]
  22. Tena-Sempere M, Barreiro ML. Leptin in male reproduction: the testis paradigm. Mol Cell Endocrinol 2002 188:9-14[CrossRef][Medline]
  23. Ojeda SR, Urbanski HF. Puberty in the rat. In: Knobil E, Neill JD (eds.), The Physiology of Reproduction, 2nd ed. New York: Raven Press; 1994: 363–410
  24. Suominen JS, Yan W, Toppari J, Kaipia A. The expression and regulation of Bcl-2 related ovarian killer (Bok) mRNA in the developing and adult rat testis. Eur J Endocrinol 2001 145:771-778[Abstract]
  25. Toppari J, Parvinen M. In vitro differentiation of rat seminiferous tubular segments from defined stages of the epithelial cycle. Morphologic and immunolocalization analysis. J Androl 1985 6:334-343[Abstract/Free Full Text]
  26. Tena-Sempere M, Pinilla L, Gonzalez LC, Dieguez C, Casanueva FF, Aguilar E. Leptin inhibits testosterone secretion from adult rat testis in vitro. J Endocrinol 1999 161:211-218[Abstract]
  27. Tena-Sempere M, Manna PR, Zhang FP, Pinilla L, Gonzalez LC, Dieguez C, Huhtaniemi I, Aguilar E. Molecular mechanisms of leptin action in rat testis: potential targets for leptin-induced inhibition of steroidogenesis and pattern of leptin receptor messenger ribonucleic acid expression. J Endocrinol 2001 170:413-423[Abstract]
  28. Chomczynski P, Sacchi N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 1987 162:156-159[Medline]
  29. Kamegai J, Wakabayashi I, Miyamoto K, Unterman TG, Kineman RD, Frohman LA. Growth hormone-dependent regulation of pituitary GH secretagogue receptor (GHS-R) mRNA levels in the spontaneous Dwarf rat. Neuroendocrinology 1998 68:312-318[CrossRef][Medline]
  30. Tamura H, Kamegai J, Sugihara H, Kineman RD, Frohman LA, Wakabayashi I. Glucocorticoids regulate pituitary growth hormone secretagogue receptor gene expression. J Neuroendocrinol 2000 12:481-485[CrossRef][Medline]
  31. Adams EF, Huang B, Buchfelder M, Howard A, Smith RG, Feighner SD, van der Ploeg LHT, Bowers CY, Fahlbusch R. Presence of growth hormone secretagogue receptor messenger ribonucleic acid in human pituitary tumors and rat GH3 cells. J Clin Endocrinol Metab 1998 83:638-642[Abstract/Free Full Text]
  32. Jeffery PL, Herington AC, Chopin LK. Expression and action of the growth hormone releasing peptide ghrelin and its receptor in prostate cancer cell lines. J Endocrinol 2002 172:R7-R11[Abstract]
  33. Bennett PA, Thomas GB, Howard AD, Feighner SD, van der Ploeg LH, Smith RG, Robinson IC. Hypothalamic growth hormone secretagogue-receptor (GHS-R) expression is regulated by growth hormone in the rat. Endocrinology 1997 138:4552-4557[Abstract/Free Full Text]
  34. Kineman RD, Kamegai J, Frohman LA. Growth hormone (GH)-releasing hormone (GHRH) and the GH secretagogue (GHS), L629,585, differentially modulate rat pituitary GHS receptor and GHRH receptor messenger ribonucleic acid levels. Endocrinology 1999 140:3581-3586[Abstract/Free Full Text]
  35. Kamegai J, Wakabayashi I, Kineman RD, Frohman LA. Growth hormone-releasing hormone receptor (GHRH-R) and growth hormone secretagogue receptor (GHS-R) mRNA levels during postnatal development in male and female rats. J Neuroendocrinol 1999 11:299-306[CrossRef][Medline]
  36. Parvinen M. Cyclic functions of Sertoli cell. In: Russell LD, Griswold MD (eds.), The Sertoli Cell. Clearwater, FL: Cache River Press; 1993: 331–347
  37. Tena-Sempere M, Huhtaniemi I. Gonadotropin receptors. In: Fauser BCJM, Rutherford AJ, Strauss JF, van Steirteghem A (eds.), Molecular Biology in Reproductive Medicine. New York: Parthenon; 1999: 165–200
  38. Murakami N, Hayashida T, Kuroiwa T, Nakahara K, Ida T, Mondal MS, Nakazato M, Kojima M, Kangawa K. Role for central ghrelin in food intake and secretion profile of stomach ghrelin in rats. J Endocrinol 2002 174:283-288[Abstract]
  39. Caminos JE, Seoane LM, Tovar SA, Casanueva FF, Dieguez C. Influence of thyroid status and growth hormone deficiency on ghrelin. Eur J Endocrinol 2002 147:159-163[Abstract]
  40. Barreiro ML, Pinilla L, Aguilar E, Tena-Sempere M. Expression and homologous regulation of GH secretagogue receptor mRNA in rat adrenal gland. Eur J Endocrinol 2002 147:677-688[Abstract]
  41. Tanaka M, Hayashida Y, Nakao N, Nakai N, Nakashima K. Testis-specific and developmentally induced expression of a ghrelin gene-derived transcript that encodes a novel polypeptide in the mouse. Biochim Biophys Acta 2001 1522:62-65[Medline]



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