Expression Analysis of <i>Gnrh1</i> and <i>Gnrhr1</i> in Spermatogenic Cells of Rat

Ciaramella, Vincenza; Chianese, Rosanna; Pariante, Paolo; Fasano, Silvia; Pierantoni, Riccardo; Meccariello, Rosaria

doi:https://doi.org/10.1155/2015/982726

International Journal of Endocrinology

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Research Article | Open Access

Volume 2015 | Article ID 982726 | https://doi.org/10.1155/2015/982726

Expression Analysis of Gnrh1 and Gnrhr1 in Spermatogenic Cells of Rat

Vincenza Ciaramella,¹Rosanna Chianese,¹Paolo Pariante,¹Silvia Fasano,¹Riccardo Pierantoni,¹and Rosaria Meccariello²

Academic Editor: Dario Acuña-Castroviejo

Received29 Nov 2014

Accepted02 Mar 2015

Published12 Mar 2015

Abstract

Hypothalamic Gonadotropin Releasing Hormone (GnRH), via GnRH receptor (GnRHR), is the main actor in the control of reproduction, in that it induces the biosynthesis and the release of pituitary gonadotropins, which in turn promote steroidogenesis and gametogenesis in both sexes. Extrabrain functions of GnRH have been extensively described in the past decades and, in males, local GnRH activity promotes the progression of spermatogenesis and sperm functions at several levels. The canonical localization of Gnrh1 and Gnrhr1 mRNA is Sertoli and Leydig cells, respectively, but ligand and receptor are also expressed in germ cells. Here, we analysed the expression rate of Gnrh1 and Gnrhr1 in rat testis (180 days old) by quantitative real-time PCR (qPCR) and by in situ hybridization we localized Gnrh1 and Gnrhr1 mRNA in different spermatogenic cells of adult animals. Our data confirm the testicular expression of Gnrh1 and of Gnrhr1 in somatic cells and provide evidence that their expression in the germinal compartment is restricted to haploid cells. In addition, not only Sertoli cells connected to spermatids in the last steps of maturation but also Leydig and peritubular myoid cells express Gnrh1.

1. Introduction

One of the most intriguing matters in reproductive endocrinology is that, in both mammalian and nonmammalian vertebrates, molecules originally identified in the brain as neuropeptides/neurohormones exert their activity in extrabrain tissues, in particular in the gonads, which express the corresponding receptors [1, 2]. Notable examples are Neuropeptide Y, Corticotropin Releasing Factor, Gonadotropin Inhibiting Factor, and Kisspeptin [1, 3, 4]. The activity of Gonadotropin Releasing Hormone (GnRH), via GnRH receptor (GnRHR), was the first to be reported in ovary and testis from fish to mammals [2, 5] as from the end of 1970s.

GnRH is the gatekeeper of reproductive functions in both sexes. This decapeptide is synthesized in the hypothalamus and is released in the median eminence to trigger the discharge of pituitary gonadotropins (Follicle Stimulating Hormone and Luteinizing Hormone (FSH and LH, resp.)). Through the main circulation, FSH and LH reach the gonads and sustain gametogenesis and steroid biosynthesis [5]. GnRH was originally isolated in the ‘70s from the hypothalamus of pig and sheep [6, 7]. Nowadays, GnRH saga comprises at least 25 GnRH molecular forms and three GnRHRs, seven-transmembrane G-coupled receptors that exhibit a wide range of subtypes [5, 8–11]. In mammals, including human, two GnRH molecular forms (GnRH1 and GnRH2) and two GnRHRs (GnRHR1 and GnRHR2) have been characterized [5, 11]. Whether GNRHR2 in humans is a functional or a remnant gene is still an unresolved matter of debate [9]. However, the main role of GnRH1 is the communication between basal hypothalamus and pituitary gland. By contrast, GnRH2, besides a supposed hypophysiotropic activity, is mainly produced in the hindbrain and exerts a neurotransmitter and/or neuromodulatory role in the control of food intake, energy balance, sexual behavior, and stress in response to many environmental cues. A third GnRH molecular form exhibiting neuromodulator activity has been only detected in the telencephalon of teleost fish. Three different GnRHRs have been detected in fish and in amphibians [5, 8, 11].

Spermatogenesis is a complex process in which mitosis, meiosis, and differentiation coexist in order to produce high quality sperm under the control of hormonal milieu. Data in mammalian and nonmammalian animal models and in cell lines have provided evidence that steroid biosynthesis and gonocyte as well as spermatogonia proliferation, spermatogenesis progression, germ cell apoptosis, sperm release, and fertilization all require the local activity of GnRH (for review see [2, 5, 12–14]). Thus, in the testis, GnRH and GnRHR are deeply involved in the autocrine and paracrine routes that modulate the communications between somatic and germ cells [2], acting in current with several local biomodulators [5, 15–21].

Foetal expression of GnRH1 and GnRHR1 has been reported during the ontogenesis of rat gonads [22] and in mouse testis high GnRH1 activity has been observed from pubertal stages to the adult, with a decline during senescence [3]. Leydig cells are the main target of GnRH activity [12, 23, 24]; GnRH1 has been detected in the interstitial fluid of rat testis [25] and competitive binding studies, immunohistochemistry, and in situ hybridization have suggested that Sertoli cells may represent the main source of GnRH (for review see [5, 12, 26]). Besides the canonical localization in somatic cells, in rat and human Gnrh1/GNRH1 and Gnrhr1/GNRHR1 transcripts have also been localized in germ cells [27]. In human, a GNRHR2 transcript is expressed in haploid postmeiotic cells and in mature sperm as well [28]. Consistently, in mouse and rat, Northern blot analysis has revealed the presence of several forms of Gnrhr1 mRNA in isolated germ cells [29]. In particular, in rat, spermatogenic cells of some seminiferous tubules also express Gnrh1 and Gnrhr1 [27]. However, at present, little is known about the localization of GnRH1 and its receptor in different spermatogenic cells. Here, we fill this gap by reporting the localization of Gnrh1 and Gnrhr1 mRNA during spermatogenesis, in adult rats, focusing on the localization of Gnrh1 and Gnrhr1 transcripts during the steps of the spermiogenesis.

2. Materials and Methods

2.1. Animals and Tissue Collection

Wistar rats (Rattus norvegicus) were housed under definite conditions (12D : 12L) and given free access to standard food and water ad libitum. Animals at 180 days postpartum (dpp) were sacrificed by decapitation under ketamine anesthesia (100 mg/kg i.p.) in accordance with local and national guidelines covering experimental animals. For each animal testes were dissected; one testis was fixed in Bouin’s fluid and processed for in situ hybridization; one was quickly frozen by immersion in liquid nitrogen and stored at −80°C until RNA extraction. Additionally, adult rat brain was also dissected, frozen, and stored at −80°C to be used as a positive control in qPCR analysis. This project was approved by the Italian Ministry of Education, University and Research (MIUR). Procedures involving animal care were conducted in accordance with the Guide for Care and Use of Laboratory Animals (National Institutes of Health guide).

2.2. Total RNA Extraction and cDNA Preparation

Total RNA was extracted from R. norvegicus tissues (testis and brain) using Trizol Reagent (1 mL/50–100 mg tissue) (Invitrogen Life Technologies, Paisley, UK) according to the manufacturer’s instructions. Total RNA was treated for 30 min at 37°C with Dnase I (10 U/sample) (Amersham Pharmacia Biotech) to eliminate any contamination of genomic DNA. RNA purity and integrity were determined by spectrophotometer analyses at 260/280 nm and by electrophoresis. Complementary DNA (cDNA) was obtained by reverse transcription using 5 μg of total RNA, 0.5 μg of oligo dT₍₁₈₎, 0.5 mM dNTP mix, 5 mM DTT, 1x first-strand buffer (Invitrogen Life Technologies), 40 U RNase Out (Invitrogen Life Technologies), and 200 U SuperScript-III RnaseH⁻ Reverse Transcriptase (Invitrogen Life Technologies) in a final volume of 20 μL, following the manufacturer’s instructions. As negative control, total RNA not treated with reverse transcriptase was used.

2.3. Cloning of Gnrh1 and Gnrhr1

To clone Gnrh1 and Gnrhr1, 1 μL of diluted (1 : 5) cDNA was used for standard PCR analysis in combination with 10 pmol of oligonucleotide primers designed on R. norvegicus nucleotide sequence (Gnrh1: 5′-agcactggtcctatgggttg-3′ and 5′-gggccagtgcatctacatct-3′; Gnrhr1: 5′-cagctttcatgatggtggtg-3′ and 5′-ttcagctgtagtttgcgtgg-3′). The predicted amplificate sizes were 221 and 370 bp, for Gnrh1 and Gnrhr1, respectively. PCR conditions were 94°C, 5 min, 1 cycle; 94°C, 30 s, 58°C, 30 s, 72°C, 45 s, 40 cycles; 72°C, 7 min. PCR products were subcloned in pGEM-T Easy Vector (Promega Corp., Madison, WI). DH5α high-efficiency competent cells were transformed and recombinant colonies were identified by blue/white colour screening. Recombinant plasmid DNA was extracted by using the QIAprep Spin Miniprep kit (Qiagen, Valencia, CA), and the insert sizes were controlled by restriction analysis with EcoRI (Fermentas, St. Leon-Rot, Germany) and then they were sequenced on both strands by Primm Sequence Service (Primm Srl, Naples Italy).

2.4. Riboprobes Synthesis and In Situ Hybridization

Specific riboprobes were synthesized by in vitro transcription using cDNA fragments of rat Gnrh1 and Gnrhr1, 221 and 370 bp, respectively, cloned previously in pGEM-T Easy Vector (Promega Corp.). The sense (control) and antisense complementary RNA (cRNA) probes were transcribed with T7 and SP6 RNA polymerases on plasmids and linearized with the appropriate restriction enzymes (SalI or NcoI) using DIG-uridine triphosphate (UTP) RNA labeling mix (Roche Diagnostics, Mannheim, Germany) as recommended by the manufacturer.

For histological observation, rat testes were fixed in Bouin’s fluid, dehydrated in ethanol, cleared in xylene, and embedded in paraffin. Tissue sections (5 μm) were stained with hematoxylin-eosin to assess sample quality. In situ hybridization was performed as follows. In brief, sections were subjected to the treatment with 10 μg/mL proteinase K (Sigma Aldrich) in 20 mM Tris-HCl; then hybridization was performed overnight at 60°C in a humidified chamber using 100 μL hybridization buffer (40% deionized formamide, 5x SSC, 1x Denhardt’s solution, 100 μg/mL sonicated salmon sperm DNA, 100 μg/mL tRNA, and 100 ng digoxigenin-labeled cRNA probe). Finally, sections were incubated for 30 min at 37°C in RNase buffer (0.5 M NaCl, 10 mM Tris-HCl, pH 7, and 5 mM EDTA) containing 20 μg/mL RNase A. Slides were observed under a light microscope (Leica LEITS DMRB; Leica Microsystem, Milan, Italy) and images were captured using a high-resolution digital camera (Leica MC 170 HD, Software Leica Application Suite 4.3).

2.5. Quantitative Real-Time PCR (qPCR)

Quantitative mRNA analysis was conducted on testis and brain to evaluate Gnrh1 and Gnrhr1 expression. All qPCRs were prepared in a final volume of 20 μL containing 1 μL of 1 : 5 diluted cDNA, 0.5 μM of each primer, and 10 μL of SSo Fast EvaGreen supermix (Bio-Rad). Assays were run twice in duplicate using the Mastercycler CFX-96 (Bio-Rad); a negative control in which cDNA was replaced by water was also included. All assays included a melting curve analysis for which all samples displayed single peaks for each primer pair. Genes of interest were normalized to the reference gene β-actin [5′-agatgacccagatcatgtttgaga-3′ and 5′-accagaggcatacagggacaa-3′; (°C) annealing 56°C; amplificate size 86 bp]; the relative quantification of the mRNA levels was performed using the comparative Cq method with the formula . Data were then reported as mean fold change ± SD over the minimal value arbitrarily assigned to a reference sample (brain sample). ANOVA followed by Duncan’s test for multigroup comparison was carried out to assess the significance of differences.

3. Results

3.1. Expression of Gnrh1 and Gnrhr1 Transcripts in R. norvegicus

Standard RT-PCR was preliminarily carried out on adult rats in order to clone Gnrh1 and Gnrhr1 cDNA fragments. Bands of the predicted size (221 and 370 bp for Gnrh1 and Gnrhr1, resp.) were observed in the testis. Therefore, the abundance of Gnrh1 and Gnrhr1 was evaluated by qPCR analysis on total RNA from adult male gonads and brain of R. norvegicus (Figures 1(a) and 1(b)). The analysis revealed the presence of the transcripts, Gnrh1 and Gnrhr1, in the testis as well as in the brain (positive control). In particular, the expression of Gnrh1 in the testis was significantly lower than in the brain () (Figure 1(a)) and that of Gnrhr1 increased 5.46-fold in the testis compared to the brain () (Figure 1(b)).

(a)

(b)

3.2. Localization of Gnrh1 and Gnrhr1 in R. norvegicus Testis

Gnrh1 and Gnrhr1 transcripts were localized by in situ hybridization in rat testis at 180 dpp (Figure 2) during the spermatogenesis. The stages were classified according to Leblond and Clermont 1952 [30].

(a)

(b)

(c)

Gnrh1 mRNA was strongly localized in the interstitial Leydig cells (LC) (Figure 2(b)) and in spermatids (SPT) in acrosome phase, Stages IX–XII (Figure 3(b)); the signal was also detected in Sertoli cells (SC) connected with SPT during the maturation phase, Stages V–VII (steps 17–19) (Figures 3(b) and 3(c)). Besides, Gnrh1 mRNA outlined each seminiferous tubule, marking peritubular myoid cells (Figures 3(b) and 3(c)).

Figure 3

Localization of Gnrh1 and Gnrhr1 by in situ hybridization in rat testis during the stage of the spermatogenetic cycle ((a)–(f)). The blue staining indicates the positive cells; few of them, representative of the different cell types, are pointed out by white arrowhead, SPT ((a)–(f)); black arrow, peritubular myoid cells ((b), (c), (e), and (f)); SC, Sertoli cells. The insets are shown in 100x magnification. Scale bars: 20 μm. The results are representative of one of three assays.

Gnrhr1 mRNA was mainly localized in SPT from Stages III to V (steps 16-17) (Figure 3(d)), when SPT are in maturation phase, until Stages XII-XIII, when SPT are in the acrosome phase (Figures 3(e) and 3(f)); a weaker signal was also observed in the interstitial compartment (LC) (Figure 2(c)).

Quiescent primary SPG (ISPG) and pachytene or leptotene spermatocytes (SPC) did not reveal any positivity. No labeling for Gnrh1 or Gnrhr1 was detected in slides incubated with sense cRNA probes (insets in Figures 2(b) and 2(c)).

A schematic representation of Gnrh1 and Gnrhr1 expression in the germinal epithelium during the spermatogenetic cycle is represented in Figure 4.

Figure 4

Schematic representation of Gnrh1 and Gnrhr1 expression in the germinal epithelium during a complete spermatogenetic cycle (modified from [45]). Columns show the sequence of germ cell generations observed in rat seminiferous tubule at that stage (I–XIV Stages). Spermatogonia: A (type A), In (intermediate), B (type B), R (resting, final DNA replication), and m (mitosis of spermatogonia). Primary spermatocytes: L (leptotene), Z (zygotene), P (pachytene), Di (diakinesis), and II (secondary spermatocytes). Phases 1–19 indicate spermatids differentiation (spermiogenesis). Horizontal lines indicate localization window of Gnrh1 and Gnrhr1, blue line and red dotted line, respectively. Vertical green lines indicate the expression window of Gnrh1 in Sertoli cells.

4. Discussion

Local activity of GnRH has been reported in male and female gonads in several animal models from molluscs to vertebrates, human included [2, 5, 13, 14, 26]. In this respect, the use of GnRH agonists and antagonists in cell lines, in vivo and in vitro, has provided evidences of GnRH involvement in the control of Leydig cells ontogenesis and activity, spermatogenesis progression, sperm release, and function ([2, 5, 13, 14, 26] and references therein).

Present data confirm earlier findings that both Gnrh1 and Gnrhr1 are expressed in rodent testis as well as in brain used as positive control. In our study total RNA was extracted from whole brain and testis and the same amount of cDNA was used for cDNA preparation. Since it is well known that in the brain Gnrh1 and Gnrhr1 are expressed in discrete brain areas and not all over the tissue [5], a dilution effect on the expression of Gnrh1 and Gnrhr1 might be postulated in the brain. Despite that, present data confirm that the brain is the major source of GnRH1. By contrast, the expression levels of Gnrhr1 here observed are higher in the testis than in the whole brain. Consistently previous results obtained by Northern blot, a less sensitive technique than qPCR, revealed that in rodents Gnrhr1 is highly expressed in the pituitary gland, the tissue mainly responsive to GnRH1, but also in whole testis and germ cells [29, 31].

By in situ hybridization we have localized Gnrh1 and Gnrhr1 inside the testis throughout the spermatogenic cycle, which in rat comprises XIV different cellular associations, with newly formed and older spermatids overlapping during the first eight stages of development [30].

In general the communications between Leydig and Sertoli cells and between Sertoli and germ cells ensure an optimal environment for the progression of germ cells and are fundamental tools to gain a successful spermatogenesis [12]. Hypogonadal (hpg) mice, a natural model with arrested reproductive development due to a congenital deficiency in GnRH synthesis leading to markedly reduced production of gonadotropins, are infertile [32]. The administration of androgens is sufficient to induce spermatogenesis in hpg mice, but the administration of recombinant human FSH, whose main targets in testis are Sertoli cells, induces the proliferation of spermatogonia and spermatogenesis progression until spermatocytes [33, 34]. Similarly, the lack of LH signalling, mainly occurring via Leydig cells, also causes spermatogenesis arrest [35, 36]. The detection of GnRH-like material in the fluid surrounding Leydig cells that express Gnrhr1 and the finding of Gnrh1 mRNA in Sertoli cells [27] suggest that GnRH1 may be produced by Sertoli cells and may act as a paracrine factor to modulate the activity of Leydig cells. Here, by in situ hybridization, we demonstrate that in adult testis Gnrh1 is expressed in Sertoli cells, in Leydig cells, and in peritubular myoid cells, confirming previous data concerning the activity of GnRH in the control of steroidogenesis and germ cells development as well as its release into the lumen of seminiferous tubules [2, 5]. Consistently, Gnrh1 mRNA has also been detected in the interstitial compartment of frog testis [15] and GnRH1 positive immunostaining has recently been reported in the interstitial compartment of mouse testis during different stages of aging, with high levels observed during reproductively active stages [3]. Interestingly, the expected expression of Gnrh1 in Sertoli cells is not widespread all over the tubules but is restricted to Stages V–VII of the spermatogenic cycle, with Sertoli cells connected to the spermatids in the last step of the maturation phase.

Present data confirm the expression of both Gnrh1 and Gnrhr1 in germ cells [27, 29], since the corresponding transcripts have been specifically localized in haploid cells but not in mitotic cells or in meiotic cells. Consistently, in mouse GnRH1 is an autocrine factor for gonocyte that expresses both ligand and receptors [3], but in the adult testis mitotic cells are devoid of any GnRH1/GnRHR1 immunoreactivity [3]. By contrast, in amphibians, which exhibit an annual reproductive cycle characterized by periods of spermatogenic arrest and periods of spermatogenesis onset and resumption, gnrh1 expression has been observed in quiescent and proliferating spermatogonia as well [15].

At the end of meiosis, haploid round spermatids undergo dynamic morphologic changes, which include acrosome formation, nuclear shaping, elongation, and tail formation in order to produce sperm cells. Hence, the functional meaning of mRNA detection in haploid cells might open several interpretations. Some mRNAs repressed in early spermatids might be recruited toward the polysomes for translation in late spermatids, whereas some other mRNAs might be transcribed in a specific time window to be immediately translated. By contrast, untranslated mRNA might be degraded at specific steps during spermiogenesis or stored in sperm in order to contribute to fertilization and embryo development, with the chromatoid body thought to be the main actor in the capture of mRNAs for degradation or storage [37]. Here we demonstrate that in rat Gnrh1 is only expressed in spermatids during the acrosome phase (Stages IX–XII), whereas Gnrhr1 is expressed in spermatids from the acrosome phase (Stages XII-XIII) throughout the maturation phase. This is consistent with the recent finding that GnRHR1 is localized in elongating spermatids in reproductively active mouse [3]. However, the expression of ligand and receptor in the same cell type during a specific time window supports the hypothesis that GnRH1 may be an autocrine modulator of acrosome biogenesis. By contrast, the detection of Gnrh1 mRNA in peritubular myoid cells and in Sertoli cells during the last step of the maturation phase (Stages V–VIII) in parallel to Gnrhr1 detection in late spermatids suggests that GnRH1 may be a paracrine modulator in sperm release and transport. Consistently, in a nonmammalian vertebrate, the anuran amphibian Pelophylax esculentus, a species that expresses two GnRH molecular forms (GnRH1 and GnRH2) and three GnRHRs (GnRHR1, R2, and R3) [38] with a functional portioning in testis [15], gnrhr2 mRNA has been localized in Sertoli cells, connected to elongating spermatids which express gnrh2 [15]. In this experimental model, buserelin, a GnRH agonist, induces spermiation [39].

GNRHR1 signaling is important for sperm binding to the human zona pellucida [40] and for the inhibition of in vivo and in vitro fertilization in rodents exerted by GnRH antagonists [41]. Furthermore, GnRH-like substances have been detected in human seminal fluid and related to the acquisition of sperm functions [42], the prostate being the major candidate for its production [43, 44].

5. Conclusions

In rat, Gnrh1 and Gnrhr1 are expressed in the testis in somatic cells and in haploid germ cells. The expression of Gnrh1/Gnrhr1 at specific steps of spermiogenesis and in Sertoli cells connected to spermatids in the late phase of maturation suggests a deep involvement in functions related to the production of high quality sperm.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgment

Financial support was provided by the Ministero Italiano dell’Istruzione, dell’Università e della Ricerca (Prin-MIUR 2010/11) to Rosaria Meccariello.

References

N. L. McGuire and G. E. Bentley, “Neuropeptides in the gonads: from evolution to pharmacology,” Frontiers in Pharmacology, vol. 1, article 114, Article ID Article 114, 2010.
View at: Publisher Site | Google Scholar
R. Meccariello, R. Chianese, T. Chioccarelli et al., “Intra-testicular signals regulate germ cell progression and production of qualitatively mature spermatozoa in vertebrates,” Frontiers in Endocrinology, vol. 5, article 69, 2014.
View at: Publisher Site | Google Scholar
S. Anjum, A. Krishna, R. Sridaran, and K. Tsutsui, “Localization of gonadotropin-releasing hormone (GnRH), gonadotropin-inhibitory hormone (GnIH), kisspeptin and GnRH receptor and their possible roles in testicular activities from birth to senescence in mice,” Journal of Experimental Zoology Part A: Ecological Genetics and Physiology, vol. 317, no. 10, pp. 630–644, 2012.
View at: Publisher Site | Google Scholar
R. Chianese, V. Ciaramella, S. Fasano, R. Pierantoni, and R. Meccariello, “Kisspeptin receptor, GPR54, as a candidate for the regulation of testicular activity in the frog Rana esculenta,” Biology of Reproduction, vol. 88, no. 3, article 73, pp. 1–11, 2013.
View at: Publisher Site | Google Scholar
R. Pierantoni, G. Cobellis, R. Meccariello, and S. Fasano, “Evolutionary aspects of cellular communication in the vertebrate hypothalamo-hypophysio-gonadal axis,” International Review of Cytology, vol. 218, pp. 69–141, 2002.
View at: Publisher Site | Google Scholar
M. Amoss, R. Burgus, R. Blackwell, W. Vale, R. Fellows, and R. Guillemin, “Purification, amino acid composition and N-terminus of the hypothalamic luteinizing hormone releasing factor (LRF) of ovine origin,” Biochemical and Biophysical Research Communications, vol. 44, no. 1, pp. 205–210, 1971.
View at: Publisher Site | Google Scholar
H. Matsuo, Y. Baba, R. M. G. Nair, A. Arimura, and A. V. Schally, “Structure of the porcine LH- and FSH-releasing hormone. I. The proposed amino acid sequence,” Biochemical and Biophysical Research Communications, vol. 43, no. 6, pp. 1334–1339, 1971.
View at: Publisher Site | Google Scholar
J. A. King and R. P. Millar, “Evolutionary aspects of gonadotropin-releasing hormone and its receptor,” Cellular and Molecular Neurobiology, vol. 15, no. 1, pp. 5–23, 1995.
View at: Publisher Site | Google Scholar
R. P. Millar, Z.-L. Lu, A. J. Pawson, C. A. Flanagan, K. Morgan, and S. R. Maudsley, “Gonadotropin-releasing hormone receptors,” Endocrine Reviews, vol. 25, no. 2, pp. 235–275, 2004.
View at: Publisher Site | Google Scholar
K. Morgan and R. P. Millar, “Evolution of GnRH ligand precursors and GnRH receptors in protochordate and vertebrate species,” General and Comparative Endocrinology, vol. 139, no. 3, pp. 191–197, 2004.
View at: Publisher Site | Google Scholar
O. Kah, C. Lethimonier, G. Somoza, L. G. Guilgur, C. Vaillant, and J. J. Lareyre, “GnRH and GnRH receptors in metazoa: a historical, comparative, and evolutive perspective,” General and Comparative Endocrinology, vol. 153, no. 1–3, pp. 346–364, 2007.
View at: Publisher Site | Google Scholar
R. M. Sharpe, “Paracrine control of the testis,” Clinics in Endocrinology and Metabolism, vol. 15, no. 1, pp. 185–207, 1986.
View at: Publisher Site | Google Scholar
R. Pierantoni, G. Cobellis, R. Meccariello et al., “Testicular gonadotropin-releasing hormone activity, progression of spermatogenesis, and sperm transport in vertebrates,” Annals of the New York Academy of Sciences, vol. 1163, pp. 279–291, 2009.
View at: Publisher Site | Google Scholar
M. Osada and N. Treen, “Molluscan GnRH associated with reproduction,” General and Comparative Endocrinology, vol. 181, no. 1, pp. 254–258, 2013.
View at: Publisher Site | Google Scholar
R. Chianese, V. Ciaramella, D. Scarpa, S. Fasano, R. Pierantoni, and R. Meccariello, “Anandamide regulates the expression of GnRH1, GnRH2, and GnRH-Rs in frog testis,” American Journal of Physiology: Endocrinology and Metabolism, vol. 303, no. 4, pp. E475–E487, 2012.
View at: Publisher Site | Google Scholar
R. Chianese, V. Ciaramella, D. Scarpa, S. Fasano, R. Pierantoni, and R. Meccariello, “Endocannabinoids and endovanilloids: a possible balance in the regulation of the testicular GnRH signalling,” International Journal of Endocrinology, vol. 2013, Article ID 904748, 9 pages, 2013.
View at: Publisher Site | Google Scholar
M.-C. Botté, Y. Lerrant, A. Lozach, A. Bérault, R. Counis, and M.-L. Kottler, “LH down-regulates gonadotropin-releasing hormone (GnRH) receptor, but not GnRH, mRNA levels in the rat testis,” Journal of Endocrinology, vol. 162, no. 3, pp. 409–415, 1999.
View at: Publisher Site | Google Scholar
J. P. Hapgood, H. Sadie, W. van Biljon, and K. Ronacher, “Regulation of expression of mammalian gonadotrophin-releasing hormone receptor genes,” Journal of Neuroendocrinology, vol. 17, no. 10, pp. 619–638, 2005.
View at: Publisher Site | Google Scholar
R. Chianese, V. Ciaramella, S. Fasano, R. Pierantoni, and R. Meccariello, “Kisspeptin drives germ cell progression in the anuran amphibian Pelophylax esculentus: a study carried out in ex vivo testes,” General and Comparative Endocrinology, vol. 211, pp. 81–91, 2015.
View at: Publisher Site | Google Scholar
S. Anjum, A. Krishna, and K. Tsutsui, “Inhibitory roles of the mammalian GnIH ortholog RFRP3 in testicular activities in adult mice,” The Journal of Endocrinology, vol. 223, no. 1, pp. 79–91, 2014.
View at: Publisher Site | Google Scholar
H. Y. Lee, M. I. Naseer, S. Y. Lee, and M. O. Kim, “Time-dependent effect of ethanol on GnRH and GnRH receptor mRNA expression in hypothalamus and testis of adult and pubertal rats,” Neuroscience Letters, vol. 471, no. 1, pp. 25–29, 2010.
View at: Publisher Site | Google Scholar
M.-C. Botté, A.-M. Chamagne, M.-C. Carré, R. Counis, and M.-L. Kottler, “Fetal expression of GnRH and GnRH receptor genes in rat testis and ovary,” Journal of Endocrinology, vol. 159, no. 1, pp. 179–189, 1998.
View at: Publisher Site | Google Scholar
A. J. Hsueh and P. B. Jones, “Extrapituitary actions of gonadotropin-releasing hormone,” Endocrine Reviews, vol. 2, no. 4, pp. 437–461, 1981.
View at: Publisher Site | Google Scholar
G. Chieffi, R. Pierantoni, and S. Fasano, “Immunoreactive GnRH in hypothalamic and extrahypothalamic areas,” International Review of Cytology, vol. 127, pp. 1–55, 1991.
View at: Publisher Site | Google Scholar
F. Petersson, G. Emons, and M. Hammar, “Testicular GnRH-receptors and direct effects of a GnRH-agonist on human testicular steroidogenesis,” Scandinavian Journal of Urology and Nephrology, vol. 23, no. 3, pp. 161–164, 1989.
View at: Publisher Site | Google Scholar
N. Ramakrishnappa, R. Rajamahendran, Y. M. Lin, and P. C. K. Leung, “GnRH in non-hypothalamic reproductive tissues,” Animal Reproduction Science, vol. 88, no. 1-2, pp. 95–113, 2005.
View at: Publisher Site | Google Scholar
J. Y. Bahk, J. S. Hyun, S. H. Chung et al., “Stage specific identification of the expression of GnRH mRNA and localization of the GnRH receptor in mature rat and adult human testis,” The Journal of Urology, vol. 154, no. 5, pp. 1958–1961, 1995.
View at: Publisher Site | Google Scholar
W. van Biljon, S. Wykes, S. Scherer, S. A. Krawetz, and J. Hapgood, “Type II gonadotropin-releasing hormone receptor transcripts in human sperm,” Biology of Reproduction, vol. 67, no. 6, pp. 1741–1749, 2002.
View at: Publisher Site | Google Scholar
P. Bull, P. Morales, C. Huyser, T. Socías, and E. A. Castellón, “Expression of GnRH receptor in mouse and rat testicular germ cells,” Molecular Human Reproduction, vol. 6, no. 7, pp. 582–585, 2000.
View at: Publisher Site | Google Scholar
C. P. Leblond and Y. Clermont, “Definition of the stages of the cycle of the seminiferous epithelium in the rat,” Annals of the New York Academy of Sciences, vol. 55, no. 4, pp. 548–573, 1952.
View at: Publisher Site | Google Scholar
S. S. Kakar, K. Grantham, L. C. Musgrove, D. Devor, J. C. Sellers, and J. D. Neill, “Rat gonadotropin-releasing hormone (GnRH) receptor: tissue expression and hormonal regulation of its mRNA,” Molecular and Cellular Endocrinology, vol. 101, no. 1-2, pp. 151–157, 1994.
View at: Publisher Site | Google Scholar
B. M. Cattanach, C. A. Iddon, H. M. Charlton, S. A. Chiappa, and G. Fink, “Gonadotrophin-releasing hormone deficiency in a mutant mouse with hypogonadism,” Nature, vol. 269, no. 5626, pp. 338–340, 1977.
View at: Publisher Site | Google Scholar
J. Singh and D. J. Handelsman, “The effects of recombinant FSH on testosterone-induced spermatogenesis in gonadotrophin-deficient (hpg) mice,” Journal of Andrology, vol. 17, no. 4, pp. 382–393, 1996.
View at: Google Scholar
H. Baines, M. O. Nwagwu, E. C. Furneaux et al., “Estrogenic induction of spermatogenesis in the hypogonadal (hpg) mouse: role of androgens,” Reproduction, vol. 130, no. 5, pp. 643–654, 2005.
View at: Publisher Site | Google Scholar
I. Huhtaniemi, P. Ahtiainen, T. Pakarainen, S. B. Rulli, F.-P. Zhang, and M. Poutanen, “Genetically modified mouse models in studies of luteinising hormone action,” Molecular and Cellular Endocrinology, vol. 252, no. 1-2, pp. 126–135, 2006.
View at: Publisher Site | Google Scholar
H. Peltoketo, F. P. Zhang, and S. B. Rulli, “Animal models for aberrations of gonadotropin action,” Reviews in Endocrine and Metabolic Disorders, vol. 12, no. 4, pp. 245–258, 2011.
View at: Publisher Site | Google Scholar
K. C. Kleene, “Patterns, mechanisms, and functions of translation regulation in mammalian spermatogenic cells,” Cytogenetic and Genome Research, vol. 103, no. 3-4, pp. 217–224, 2003.
View at: Publisher Site | Google Scholar
R. Chianese, V. Ciaramella, S. Fasano, R. Pierantoni, and R. Meccariello, “Anandamide modulates the expression of GnRH-II and GnRHRs in frog, Rana esculenta, diencephalon,” General and Comparative Endocrinology, vol. 173, no. 3, pp. 389–395, 2011.
View at: Publisher Site | Google Scholar
S. Minucci, L. Di Matteo, G. C. Baccari, and R. Pierantoni, “A gonadotropin releasing hormone analog induces spermiation in intact and hypophysectomized frogs, Rana esculenta,” Experientia, vol. 45, no. 11-12, pp. 1118–1121, 1989.
View at: Publisher Site | Google Scholar
P. Morales, E. Pizarro, M. Kong, and C. Pasten, “Sperm binding to the human zona pellucida and calcium influx in response to GnRH and progesterone,” Andrologia, vol. 34, no. 5, pp. 301–307, 2002.
View at: Publisher Site | Google Scholar
P. Morales, C. Pasten, and E. Pizarro, “Inhibition of in vivo and in vitro fertilization in rodents by gonadotropin-releasing hormone antagonists,” Biology of Reproduction, vol. 67, no. 4, pp. 1360–1365, 2002.
View at: Publisher Site | Google Scholar
S. I. Izumi, T. Makino, and R. Iizuka, “Immunoreactive luteinizing hormone-releasing hormone in the seminal plasma and human semen parameters,” Fertility and Sterility, vol. 43, no. 4, pp. 617–620, 1985.
View at: Google Scholar
N. Azad, S. Uddin, N. La Paglia et al., “Luteinizing hormone-releasing hormone (LHRH) in rat prostate: characterization of LHRH peptide, messenger ribonucleic acid expression, and molecular processing of LHRH in intact and castrated male ratss,” Endocrinology, vol. 133, no. 3, pp. 1252–1257, 1993.
View at: Google Scholar
R. B. White, J. A. Eisen, T. L. Kasten, and R. D. Fernald, “Second gene for gonadotropin-releasing hormone in humans,” Proceedings of the National Academy of Sciences of the United States of America, vol. 95, no. 1, pp. 305–309, 1998.
View at: Publisher Site | Google Scholar
B. Perey, Y. Clermont, and C. P. Leblond, “The wave of the seminiferous epithelium in the rat,” The American Journal of Anatomy, vol. 108, no. 1, pp. 47–77, 1961.
View at: Publisher Site | Google Scholar

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Copyright © 2015 Vincenza Ciaramella et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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