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Differential Gonadotropin-Releasing Hormone (GnRH) and GnRH Receptor Messenger Ribonucleic Acid Expression Patterns in Different Tissues of
     Department of Neurobiology, Weizmann Institute of Science (T.D.S.-H., N.B.-A., Y.K.), Rehovot 76100, Israel; and Department of Chemistry and Bioscience, Chalmers University of Technology (T.B.), 405 30 Gothenburg, Sweden

    Address all correspondence and requests for reprints to: Dr. Yitzhak Koch, Department of Neurobiology, Weizmann Institute of Science, Rehovot 76100, Israel. E-mail: y.koch@weizmann.ac.il.

    Abstract

    GnRH, the main regulator of reproduction, is produced in a variety of tissues outside of the hypothalamus, its main site of synthesis and release. We aimed to determine whether GnRH produced in the female rat pituitary and ovaries is involved in the processes leading to ovulation. We studied the expression patterns of GnRH and GnRH receptor (GnRH-R) in the same animals throughout the estrous cycle using real-time PCR. Hypothalamic levels of GnRH mRNA were highest at 1700 h on proestrus, preceding the preovulatory LH surge. No significant changes in the level of hypothalamic GnRH-R mRNA were detected, although fluctuations during the day of proestrus are evident. High pituitary GnRH mRNA was detected during the day of estrus, in the morning of diestrus 1, and at noon on proestrus. Pituitary GnRH-R displayed a similar pattern of expression, except on estrus, when its mRNA levels declined. Ovarian GnRH mRNA levels increased in the morning of diestrus 1 and early afternoon of proestrus. Here, too, GnRH-R displayed a somewhat similar pattern of expression to that of its ligand. To the best of our knowledge, this is the first demonstration of a GnRH expression pattern in the pituitary and ovary of any species. The different timings of the GnRH peaks in the three tissues imply differential tissue-specific regulation. We believe that the GnRH produced in the anterior pituitary and ovary could play a physiological role in the preparation of these organs for the midcycle gonadotropin surge and ovulation, respectively, possibly via local GnRH-gonadotropin axes.

    Introduction

    THE HYPOTHALAMIC GnRH is the main regulator of reproductive functions in all vertebrate species. This decapeptide is released from nerve endings of hypothalamic neurons into the portal system, leading it into the anterior pituitary, where it binds to specific G protein-coupled receptors on gonadotrope cell membranes and induces the synthesis and release of LH and FSH into the systemic circulation.

    In both humans and rats, GnRH and its receptor have been found to be synthesized in a myriad of tissues outside of their classical sites of production, such as pituitary (1, 2, 3, 4, 5), ovary (6, 7, 8, 9, 10), endometrium and placenta (11, 12), breast tissue and mammary glands (13, 14, 15, 16), liver, heart, skeletal muscles, kidney (17), spleen lymphocytes (18), and T cells (19). GnRH and GnRH receptor (GnRH-R) production in the various extrahypothalamic organs and glands has not yet been quantified, and their roles in these tissues have not been elucidated. We hypothesize, however, that GnRH carries out local functions in extrahypothalamic tissues, which it performs via autocrine/paracrine mechanisms. In our studies of the role of GnRH in extrahypothalamic organs, we chose to focus on the pituitary and ovary.

    Evidence has been presented showing that GnRH is produced in and released by gonadotrope cells (20) or gonadotrope and corticotrope cells (3) of rat pituitaries. Moreover, in vitro findings point to a potential role for the endogenous pituitary GnRH in the maintenance of basal LH levels (20, 21, 22). Indeed, our research group has shown that the addition of a selective GnRH antagonist, antide, to primary pituitary cell cultures causes a stark reduction in the basal secretion of LH (21).

    In the rat ovary, GnRH and its receptor are produced by granulosa cells (7, 8, 9, 10). A myriad of functions have been suggested for GnRH in this organ (recently reviewed in Ref. 23), including a role in oocyte maturation (24) and follicular atresia or selection (8), an effect on the corpus luteum (25), as well as an effect on the fertilization process (26). Interestingly, it has been shown that GnRH can induce ovulation in hypophysectomized rats (27, 28, 29). Importantly, unlike the pituitary GnRH-R, ovarian receptors are most likely activated by locally produced GnRH, because this peptide is found in only minute, undetectable amounts in the peripheral circulation (30). Hence, it is of relevance to study changes in locally produced ovarian GnRH throughout the estrous cycle.

    The chain of hormonal and neuronal events that lead to the preovulatory gonadotropin surge, the pivotal event in the mammalian reproductive cycle, has been studied intensively, although it is still not completely understood (for a concise review, see Ref. 31). We raise the possibility that local GnRH, produced by pituitary and ovarian cells, could be one of the missing links in this complex chain of events leading to a midcycle gonadotropin surge and subsequent ovulation. In the present study we hypothesized that if GnRH produced locally in the ovary and/or pituitary is involved in the processes leading to ovulation, we might be able to discern periovulatory changes in its expression. Thus, our main venue of research consisted of investigating the pattern of GnRH and GnRH-R expression in the pituitary and ovary of the female rat throughout the estrous cycle compared with the pattern observed in the hypothalamus. We focused on the anticipated time of the proestrous LH surge to test the possible involvement of the locally produced pituitary and ovarian GnRH in this event. mRNA levels were measured relative to stably expressed genes using real-time PCR. We found fluctuations in the levels of GnRH and GnRH-R mRNA in all three tissues throughout the estrous cycle. We therefore suggest that locally produced ovarian and pituitary GnRH might fulfill regulatory roles in the processes leading to ovulation.

    Materials and Methods

    Animals: estrous cycle study

    All animals were purchased from Harlan Laboratories (Rehovot, Israel), and all experiments were carried out in compliance with the regulations of the Weizmann Institute of Science using accepted standards of humane animal care. Intact female Wistar rats (7–9 wk old) were used in this study. Animals were housed under constant conditions of temperature and humidity, with lights on between 0600–2000 h and food available ad libitum. Estrous cycles were monitored via vaginal smears, and only rats showing at least three consecutive 4-d cycles were used.

    Animals were killed by decapitation at the following times: at 1000 and 1600 h on diestrus 1 (n = 12 and 4) and diestrus 2 (n = 10 and 4); on proestrus at 0900 h (n = 4), 1200 h (n = 10), 1400 h (n = 4), 1500 h (n = 4), 1600 h (n = 5), 1700 (n = 8), 1900 h (n = 10), and 2130 h (n = 6); and on estrus at 0900 h (n = 7) and at 1400 h (n = 5). Trunk blood was collected, and serum was separated and frozen until subsequent quantitative determination by RIA for LH. Tissues were immediately removed and placed in 10 vol RNA Later (Ambion, Inc., Austin, TX) until subsequent RNA extraction. The hypothalamus was dissected out to a depth of approximately 3 mm with the following borders: the anterior edge of the optic chiasm, the anterior edge of the mammillary bodies, and the two hypothalamic sulci on either lateral side. Both ovaries were removed, and oviducts were examined for the presence of ova only in the groups killed on the morning of estrus. The anterior pituitary was removed together with the attached posterior lobe.

    Animals: quantitation of GnRH peptide in various tissues

    Six- to 8-wk-old female Wistar rats were killed, and hypothalami, ovaries, and anterior pituitaries were removed as described above. Tissues were pooled (for details, see Fig. 1), boiled, homogenized, and boiled again (32). Homogenates were centrifuged, and the GnRH concentration in the resulting supernatant was determined as described previously (33).

    FIG. 1. GnRH content in the hypothalamus, pituitary, and ovary of cycling female rats (n = 6). Tissues were removed, pooled (two pools of three hypothalami each; two pools containing three pairs of ovaries each; one pool containing all six pituitaries), and homogenized, and GnRH was quantitated using RIA. The GnRH content (average of two pools for hypothalamus and ovary, result of a single pituitary pool) ± SEM are presented on a logarithmic scale. Numbers above each column represent the (average) amount of GnRH, in picograms per organ, found in each tissue. For details, see Materials and Methods.

    RIA for serum LH and for GnRH content in different tissues

    The exact procedures employed for the quantification of LH and GnRH using RIA were described previously (21, 34). Serum LH levels were determined using the kit for rat LH provided by the National Institute of Arthritis, Metabolism and Digestive Diseases, Rat Pituitary Program. LH levels are expressed in terms of the RP-3 reference preparation. The intraassay coefficients of variation for LH and GnRH RIA were 2% and 3.2%, respectively, and the assay sensitivity was 0.2 ng/tube for LH and 3.5 pg/tube for GnRH.

    RNA purification

    Tissues were removed from the RNA Later, weighed, and homogenized in 0.5 ml Tri-Reagent (Molecular Research Center, Inc., Cincinnati, OH). After the addition of 100 μl chloroform and phase separation by centrifugation (at 4 C), the aqueous layer was washed with an equal volume of 70% ethanol and loaded onto an RNeasy minicolumn (Qiagen, Hilden, Germany). The procedures for RNA isolation and purification as well as on-column deoxyribonuclease treatment (Qiagen) were then carried out as detailed in the manufacturer’s instructions. RNA samples were eluted in nuclease-free water (Qiagen). The RNA concentration was quantified using a NanoDrop machine (NanoDrop Technologies, Wilmington, DE), and its RNA purity was assessed on the same machine using 260:280 and 260:230 nM ratios. All samples had 260:280 nM ratios between 1.8 and 2.1, and 260:230 nM ratios above 1.7. RNA integrity was assessed by examining the 28S and 18S bands of representative samples loaded onto a 1.5% agarose gel stained with ethidium bromide.

    RT

    For each tissue, equal amounts of all RNA samples were reverse transcribed simultaneously. Ovarian and hypothalamic RNA samples (2 μg each) were reverse transcribed using Moloney murine leukemia virus reverse transcriptase ribonuclease H+ (Promega Corp., Madison, WI) according to the manufacturer’s instructions. Each reaction contained 0.5 μg oligo(deoxythymidine) (Amersham Biosciences, Piscataway, NJ), 0.52 mM of each deoxy-NTP (MBI Fermentas, St. Leon-Rot, Germany), 25 U RNAguard ribonuclease inhibitor (Amersham Biosciences), 5 μl of the 5x Moloney murine leukemia virus RT reaction buffer (Promega Corp.), and 200 U of the enzyme in a total volume of 25 μl. Pituitary RNA samples (4 μg each) were reverse transcribed using the SuperScript II ribonuclease H– reverse transcriptase kit (Invitrogen Life Technologies, Inc., Carlsbad, CA). Each 20-μl reaction contained 0.5 μg oligo(deoxythymidine) (Amersham Biosciences), 0.5 mM of each deoxy-NTP (MBI Fermentas), 40 U porcine liver ribonuclease-inhibitor (Takara Bio, Inc., Shiga, Japan), 2 μl 0.1 M dithiothreitol, 4 μl of the 5x First-Strand Buffer (Invitrogen Life Technologies, Inc.), and 40 U of the enzyme.

    All RT reactions were performed at 42 C and contained a negative control, which consisted of nuclease-free water instead of RNA. The linearity of the RT reaction was evaluated using triplicate serial dilutions of an RNA pool, reverse transcribed as detailed above, and assayed in the real-time PCR for two genes: GnRH and cyclophillin. The reaction efficiencies (E = 97% and 100%, respectively) and correlation coefficients (r2 = 0.986 and 0.995, respectively), derived from the RNA dilution series, indicated that the RT reaction was linear under the conditions used. Several RT reactions contained duplicate RNA samples (n = 23 duplicates) to assess the RT-PCR variability.

    Gene-specific primers and TaqMan hybridization probes

    Primers were designed on two different exons so as to span one intronic sequence. TaqMan hybridization probes were designed to span an exon-exon junction (TIB-Molbiol, Berlin, Germany). All primer and probe sequences, PCR product sizes, and annealing temperatures used are listed in Table 1. To verify the identities of the PCR products obtained using each primer combination, each product was loaded onto an ethidium bromide-stained agarose gel, and the resulting band was purified and sequenced using the ABI Gene Scanner and the ABI BigDye Terminator Cycle Sequencing Kit (PerkinElmer, Applied Biosystems, Foster City, CA).

    TABLE 1. Primer and probe sequences and optimized reaction conditions

    Relative real-time PCR

    All real-time PCRs were carried out on a Rotor-Gene 3000 (Corbett Research, Sydney, Australia), using the Absolute QPCR Master Mix (ABgene, Surrey, UK) with or without SYBR Green-I, according to the assay type. Reaction protocols had the following format: 15 min at 95 C for enzyme activation, followed by 40–50 cycles of: 15 sec at 95 C, 30 sec at the appropriate annealing temperature (see Table 1), and 15 sec at 72 C, at the end of which fluorescence was measured with the Rotor-Gene. SYBR Green-I assays also included a melt curve at the end of the cycling protocol, with continuous fluorescence measurement from 65–99 C. All reactions contained the same amount of cDNA, 10 μl Absolute QPCR Master Mix, primers or primers and probe diluted according to an optimized combination (see Table 1), and UltraPure PCR-grade water (Fisher Biotec, Subiaco, Australia) to a final volume of 20 μl.

    Each real-time PCR included a no-template control as well as five or six serial 4-fold dilutions, in duplicate, of a cDNA pool containing all experimental samples of the respective tissue. The prenormalized DNA quantity of each gene in every sample was estimated relative to this dilution series. This dilution series also served to assess the reaction performance (E and r2). The threshold cycle (Ct) was set so as to obtain the highest reaction efficiency and correlation coefficient. Because not all samples of a particular tissue (n = 93) could be assayed simultaneously for each gene, a common set of at least five samples was included in every reaction for interreaction calibration.

    Preliminary validation of reference genes

    A common practice to compensate for differences in the steps preceding the PCR is normalization of gene expression to reference genes. For proper normalization, the expression of reference genes should not vary across the experimental conditions. A panel of four candidate reference genes was therefore tested in all experimental samples (n = 93/tissue) to identify the most stably expressed genes at all tested times. Primer sequences, accession numbers, and reaction conditions for all tested reference genes are listed in Table 1 (see gene names followed by an asterisk). The two most suitable reference genes in each tissue were identified by three approaches.

    The first approach, developed and described by Vandesompele et al. (35), identifies the pair of genes whose DNA quantities fluctuate the least relative to each other. It is based on the assumption that the expression ratio of two ideal reference genes that are not coregulated, is constant across experimental conditions. This approach is implemented in the freely available Excel macro, GeNorm (35).

    The second approach, developed and described by Andersen et al. (36), is based on the decomposition of expression values of each candidate reference gene into technical and biological constituents. The stability of each gene across all samples is determined both within and between groups (e.g. time groups), and the most stable single gene and pair of genes are identified. This approach is implemented in a freely available Excel Add-in, NormFinder (36).

    The most obvious advantage of NormFinder is that it examines the expression stability of each single candidate gene independently and not in relation to the other genes, as GeNorm does. This is important in light of our limited knowledge regarding coregulation. Moreover, NormFinder also tests for combinations of genes that may compensate for each other’s fluctuations. This is helpful in situations where none of the candidate reference genes is found to be stably expressed. The disadvantage of NormFinder, which is more critical when a small number of genes are tested, lies in the possibility that several of the candidate genes display similar expression trends or fluctuations in the experimental data. This causes skewing of the average intergroup variation, which should normally be close to zero for a proper identification of the least variable gene. Although the tendency of GeNorm to select coregulated genes increases with the number of tested genes, NormFinder will have more chances of selecting a stably expressed gene or a combination of two genes whose fluctuations compensate one another.

    The third approach, which we term least absolute variation, is based on the low probability that biological variability in gene expression will be precisely countermatched by (opposite) technical variability, yielding low variance in prenormalized DNA amounts of a large set of samples. According to this approach, low intersample variability of prenormalized DNA amounts, assessed by the spread, i.e. the SD, of PCR Ct indicates both biological and technical similarities. To determine whether the obtained spread in Ct values of all samples assayed for each candidate reference gene is indeed small, we compared it to the reproducibility of RT-PCRs. Unlike GeNorm and NormFinder, this approach provides information regarding the degree of stability (magnitude of the SD) of each gene.

    Data analysis and statistics

    The relative amount of GnRH or GnRH-R mRNA in each sample was calculated by dividing the prenormalized DNA quantity of these genes (calibrated to account for interreaction differences), obtained from the dilution series (see above), by the geometric mean of the DNA quantities of the two most suitable reference genes (35). This normalized DNA quantity is hereafter referred to as the relative expression level of GnRH or GnRH-R.

    All statistical analyses were performed using JMP IN Statistical Discovery software (version 5.1, SAS Institute, Inc., Cary, NC). For each tissue and gene, the distribution of the obtained LH levels and relative amounts of GnRH and GnRH-R in all samples were tested for normality using the Kolmogorov-Smirnov-Lilliefor test. In all cases studied, the log-transformed data displayed a normal distribution (P > 0.05). Statistical evaluation of differences between time groups was performed using both parametric and nonparametric multiple comparison tests (one-way ANOVA and the Wilcoxon rank-sum test, respectively), followed by pairwise comparisons of means using the least significant difference test and Student’s t test at a confidence level of 95%.

    Results

    Quantitation of GnRH content in different tissues

    The amount of GnRH peptide found in the ovary, pituitary, and hypothalamus of 8-wk-old rats is depicted in Fig. 1. Compared with the hypothalamus, we found about 430-fold less GnRH in the pituitary and about 170-fold less GnRH in the ovary.

    Selection of suitable reference genes

    Table 2 indicates, for each tissue, the two most suitable reference genes found by each of the three methods employed. The variability of RT-PCRs was estimated from RNA samples reverse transcribed in duplicate and assayed together in the real-time PCR. It was measured as the SD of duplicate reactions’ Ct values and found to be, on the average, SD = 0.22 (range, 0.001–0.42; n = 23). This corresponds well to the RT-PCR reproducibility estimate recently published by Stahlberg et al. (37): SDmRNA = 0.11–0.60. The SD of Ct values obtained for the best candidate reference genes in all tissue was found to range between 0.30 and 0.62 (Table 2), which overlaps the RT-PCR reproducibility values mentioned above. Therefore, we chose the two most suitable reference genes for each tissue to be those with the lowest SD. Thus, in pituitary and hypothalamic samples, the geometric average of cyclophillin and RPL19 expression was used to normalize GnRH and GnRH-R expression; in ovarian samples, the geometric average of cyclophillin and ?-actin expression was used to normalize GnRH and GnRH-R expression.

    TABLE 2. The most stably expressed reference genes per tissue

    Relative levels of GnRH expression during the rat estrous cycle

    In our colony, the preovulatory LH surge was found to occur on proestrus at around 1900 h (Fig. 2A), when serum LH levels were 29.1 ± 1.75 ng/ml (mean ± SEM). Fluctuations in the relative expression levels of GnRH during the estrous cycle were found in all three tissues (by one-way ANOVA, P < 0.05; by Wilcoxon rank-sums test, P < 0.05 for all three tissues). In the hypothalamus (Fig. 2B), the highest GnRH expression was found on proestrus at 1700 h, whereas the lowest expression level was found on this day at 1900 h.

    FIG. 2. The expression pattern of GnRH mRNA in the hypothalamus, pituitary, and ovary throughout the estrous cycle of female rats in relation to serum LH levels. Serum LH levels (A) and GnRH expression patterns in the hypothalamus (B), pituitary (C), and ovary (D) are shown. Numbers on the abscissa represent hours of the respective estrous cycle days. Values are the mean ± SEM of four to 11 rats per time group. LH levels are expressed in terms of the RP-3 reference preparation, in nanograms per milliliter of serum. Relative GnRH mRNA expression levels were calculated by dividing the quantity of GnRH (derived from the cDNA dilution curve) by the geometric average of the quantities of the two most suitable reference genes (cyclophillin and RPL19 in the hypothalamus and pituitary, cyclophillin and ?-actin in the ovary; see Table 2). Different letters indicate statistically significant differences between groups (P < 0.05). For details, see Materials and Methods.

    In the pituitary (Fig. 2C), the highest relative GnRH expression was detected on the morning and early afternoon of estrus. Peak proestrous amounts of GnRH mRNA were noted at noon and 1700 h. On diestrus 1 and 2, morning levels were higher than afternoon levels.

    In the ovary (Fig. 2D), peak GnRH expression levels occurred at 1000 h on diestrus 1 and 1400 h on proestrus, whereas the lowest expression levels occurred in the late afternoon-evening of proestrus and on estrus.

    Relative levels of GnRH-R expression during the rat estrous cycle

    Significant variations in the level of GnRH-R mRNA during the estrous cycle were detected in the pituitary (by one-way ANOVA, P < 0.001; by Wilcoxon rank-sums test, P < 0.0001) and ovary (by one-way ANOVA, P < 0.05; by Wilcoxon rank-sums test, P < 0.005). The hypothalamic GnRH-R mRNA (Fig. 3A) displayed only nonsignificant changes during the estrous cycle, possibly due to relatively high interindividual variability.

    FIG. 3. The expression pattern of GnRH-R mRNA in the hypothalamus, pituitary, and ovary throughout the estrous cycle of female rats. GnRH-R expression patterns in the hypothalamus (A), pituitary (B), and ovary (C) are shown. Values are the mean ± SEM of four to 11 rats per time group. Numbers on the abscissa represent hours of the respective estrous cycle days. Relative GnRH-R mRNA expression levels were calculated as described in Fig. 2. Different letters indicate statistically significant differences between groups (P < 0.05). For details, see Materials and Methods.

    In the pituitary gland (Fig. 3B), the highest relative amounts of GnRH-R mRNA were detected in the mornings of diestrus 1 and 2, followed by a sharp reduction in the afternoon of these days. On proestrus, pituitary GnRH-R expression was low in the morning and rose back to diestrous 1 and 2 (morning) levels at noon, followed by significant transient drops at 1400 and 1600 h and a second peak at 1700 h. Relatively low levels of GnRH-R expression were measured on estrus in this tissue.

    In the ovary, GnRH-R mRNA expression reached peak levels at 1400 h on proestrus, followed by a second lower peak at 1700 h (Fig. 3C). The lowest expression levels occurred at 1400 h on estrus and noon on proestrus.

    Discussion

    In this study, GnRH and GnRH-R expression levels during the estrous cycle were measured using real-time RT-PCR. Reliable use of this technology requires that errors in quantification that might occur at various stages of the experimental procedure (e.g. RNA extraction, RT, etc.) be controlled. The most common method employed to this end consists of normalizing the expression of the gene of interest by that of one, or preferably more, reference genes, whose expression level is invariant across the relevant experimental conditions. Although several tools to identify the most stably expressed genes in a set of experimental samples have been developed (35, 36, 38, 39), none is, as yet, widely accepted and used. Therefore, we chose to apply three different approaches (described in Materials and Methods) to identify the two most suitable reference genes in each of the three tissues studied.

    We obtained extremely low intersample Ct variability for our candidate reference genes (see SD values in Table 2), which is within the reproducibility range of RT-PCRs (37). This might be a somewhat rare situation, reflecting extremely small technical and biological errors. Nonetheless, we believe that under such circumstances, measurements of variation in prenormalized DNA quantities provides the most direct means of assessing the stability of candidate reference genes. Therefore, we chose the two most suitable reference genes to be those with the lowest SD. We believe that these reference genes could be used in future studies employing similar experimental systems.

    When comparing the amounts of GnRH produced by the pituitary or ovary to those found in the hypothalamus, one should take into consideration the different modes of action that this peptide undertakes in the different tissues. Thus, although GnRH functions as an endocrine hormone once it is released from the hypothalamus into the portal system, in extrahypothalamic regions it probably plays an autocrine/paracrine role. One might therefore expect lower levels of GnRH production in extrahypothalamic tissues compared with the hypothalamic levels.

    The present study is the first demonstration of a GnRH mRNA expression pattern in the pituitary and ovary throughout the estrous cycle of any species, whereas the pattern of hypothalamic GnRH expression has been studied previously (40, 41, 42, 43, 44). Our findings corroborate with previous demonstrations of a peak in hypothalamic GnRH mRNA levels in the later afternoon-early evening hours of proestrus, slightly before the gonadotropin surge (42, 43, 44). Similarly, peak levels of GnRH peptide have been found in the mediobasal hypothalamus (45, 46) and in the portal blood (47) on the evening of proestrus. This increase probably serves either to increase the availability of hypothalamic GnRH in preparation for the imminent GnRH surge or to replenish this peptide’ stores that were depleted by the surge.

    Expression of the receptor for GnRH in the mediobasal hypothalamus was previously shown to be modulated during the rat estrous cycle (48), such that peak levels were observed on the morning of proestrus and the evening of estrus. It is noteworthy that very few times points of the estrous cycle were sampled in that study. In our investigation, the entire hypothalamus was analyzed rather than specific nuclei, and only nonsignificant fluctuations in the expression of GnRH-R during the estrous cycle were observed. It is possible that the GnRH-R are differentially modulated at various hypothalamic sublocations. The hypothalamic GnRH-R are probably involved in autoregulatory feedback mechanisms in this tissue.

    In the pituitary, the rise in GnRH mRNA we observed at noon on proestrus is interesting in light of the increase in LH mRNA that begins more or less at the same time (49). It is possible that the local pituitary GnRH plays a role in the regulation of LH production, because at this time, GnRH levels in the median eminence (50) and portal system (47) are still low. As mentioned previously, a potential role for pituitary GnRH in the release of LH has been demonstrated in vitro (21). The physiological significance of the high estrous levels of GnRH mRNA in the pituitary is unclear at this point. It might be noteworthy that FSH mRNA has also been shown to increase on the day of estrus (51).

    Interestingly, there appears to be a temporal correlation between the expression pattern of GnRH and that of its receptor in the pituitary throughout the estrous cycle, except on estrus. It is possible that endogenous pituitary GnRH participates in the regulation of its receptor in this gland or that the two are coregulated. Pituitary GnRH-R expression and content as well as GnRH binding to pituitary receptors during the rat estrous cycle have been investigated in the past, although results are somewhat contradictory (52, 53, 54, 55, 56). Nevertheless, a heightened GnRH-binding capacity of the pituitary during the day or so before the preovulatory gonadotropin surge appears to be a result of increased receptor synthesis. This increase, which might be generated by locally produced pituitary GnRH, is postulated to induce heightened pituitary responsiveness to hypothalamic GnRH stimulation (57).

    In the ovary, we observed relatively high GnRH and GnRH-R expression in the early afternoon of proestrus and high GnRH-R mRNA levels also around the time of the preovulatory surge. In an earlier investigation (58), it was reported that cyclical changes during the estrous cycle in the expression of the ovarian GnRH-R are specific to the stage of follicular development, such that they were observed only in corpora lutea and atretic follicles. In these two types of follicles, peak GnRH-R levels were observed in the evening of proestrus, with a second increase in the morning of estrus observed only in atretic follicles. The researchers suggested that ovarian GnRH might be involved in follicular atresia and possibly also in the induction of ovulation. The fact that we detected peak GnRH expression concomitantly with peak GnRH receptor expression in the ovary is intriguing and raises the possibility that here too, GnRH regulates the expression of its own receptor or that the two are coregulated.

    An interesting observation was recently published (59), suggesting that oocytes of the gilthead sea bream produce and release gonadotropins, and that this release can be enhanced by a GnRH analog or reduced by a GnRH antagonist. As the researchers point out, this discovery raises the interesting possibility of a local GnRH-gonadotropin axis within the fish ovary. In preliminary RT-PCR experiments (performed in collaboration with the laboratory of N. Dekel at the Weizmann Institute), we also identified LH? expression in rat and mouse oocytes. One could thus envisage local GnRH-gonadotropin axes within the pituitary and ovaries of mammalian females. Such local regulatory axes could contribute to or finely tune the hypothalamic-pituitary-gonadal axis, for instance by priming the relevant organs in preparation for the preovulatory peak (in the case of the pituitary) and ovulation (in the case of the ovaries). Nonetheless, research in this direction has yet to be conducted.

    The present report presents a detailed and precise pattern of GnRH and GnRH-R expression during the rat estrous cycle. We propose that in the adult female rat, the production of GnRH and GnRH-R is locally regulated in the pituitary and ovary, in accordance with the animal’s reproductive state or phase of the estrous cycle. The earlier increase in GnRH production in the pituitary and ovary compared with the hypothalamus during the proestrous stage of the sexual cycle might indicate that this peptide participates in the preparation of these organs for the imminent preovulatory surge, possibly via local GnRH-gonadotropin axes. One should not preclude, however, nonreproductive autocrine/paracrine roles of GnRH in extrahypothalamic tissues.

    References

    Bauer TW, Moriarty CM, Childs GV 1981 Studies of immunoreactive gonadrotropin releasing hormone (GnRH) in the rat anterior pituitary. J Histochem Cytochem 29:1171–1178

    Li JY, Knapp RJ, Sternberger LA 1984 Immunocytochemistry of a "private" luteinizing-hormone-releasing hormone system in the pituitary. Cell Tissue Res 235:263–266

    May V, Wilber JF, U’Prichard DC, Childs GV 1987 Persistence of immunoreactive TRH and GnRH in long-term primary anterior pituitary cultures. Peptides 8:543–558

    Pagesy P, Li JY, Berthet M, Peillon F 1992 Evidence of gonadotropin-releasing hormone mRNA in the rat anterior pituitary. Mol Endocrinol 6:523–528

    Miller GM, Alexander JM, Klibanski A 1996 Gonadotropin-releasing hormone messenger RNA expression in gonadotroph tumors and normal human pituitary. J Clin Endocrinol Metab 81:80–83

    Aten RF, Polan ML, Bayless R, Behrman HR 1987 A gonadotropin-releasing hormone (GnRH)-like protein in human ovaries: similarity to the GnRH-like ovarian protein of the rat. J Clin Endocrinol Metab 64:1288–1293

    Jones PB, Conn PM, Marian J, Hsueh AJ 1980 Binding of gonadotropin releasing hormone agonist to rat ovarian granulosa cells. Life Sci 27:2125–2132

    Whitelaw PF, Eidne KA, Sellar R, Smyth CD, Hillier SG 1995 Gonadotropin-releasing hormone receptor messenger ribonucleic acid expression in rat ovary. Endocrinology 136:172–179

    Oikawa M, Dargan C, Ny T, Hsueh AJ 1990 Expression of gonadotropin-releasing hormone and prothymosin- messenger ribonucleic acid in the ovary. Endocrinology 127:2350–2356

    Clayton RN, Eccleston L, Gossard F, Thalbard JC, Morel G 1992 Rat granulosa cells express the gonadotrophin-releasing hormone gene: evidence from in-situ hybridization histochemistry. J Mol Endocrinol 9:189–195

    Khodr GS, Siler-Khodr T 1978 Localization of luteinizing hormone-releasing factor in the human placenta. Fertil Steril 29:523–526

    Seeburg PH, Adelman JP 1984 Characterization of cDNA for precursor of human luteinizing hormone releasing hormone. Nature 311:666–668

    Amarant T, Fridkin M, Koch Y 1982 Luteinizing hormone-releasing hormone and thyrotropin-releasing hormone in human and bovine milk. Eur J Biochem 127:647–650

    Baram T, Koch Y, Hazum E, Fridkin M 1977 Gonadotropin-releasing hormone in milk. Science 198:300–302

    Levi LN, Ben-Aroya N, Tel-Or S, Palmon A, Burstein Y, Koch Y 1996 Expression of the gene for the receptor of gonadotropin-releasing hormone in the rat mammary gland. FEBS Lett 379:186–190

    Palmon A, Ben Aroya N, Tel-Or S, Burstein Y, Fridkin M, Koch Y 1994 The gene for the neuropeptide gonadotropin-releasing hormone is expressed in the mammary gland of lactating rats. Proc Natl Acad Sci USA 91:4994–4996

    Kakar SS, Jennes L 1995 Expression of gonadotropin-releasing hormone and gonadotropin-releasing hormone receptor mRNAs in various non-reproductive human tissues. Cancer Lett 98:57–62

    Azad N, Emanuele NV, Halloran MM, Tentler J, Kelley MR 1991 Presence of luteinizing hormone-releasing hormone (LHRH) mRNA in rat spleen lymphocytes. Endocrinology 128:1679–1681

    Chen A, Ganor Y, Rahimipour S, Ben-Aroya N, Koch Y, Levite M 2002 The neuropeptides GnRH-II and GnRH-I are produced by human T cells and trigger laminin receptor gene expression, adhesion, chemotaxis and homing to specific organs. Nat Med 8:1421–1426

    Krsmanovic LZ, Martinez-Fuentes AJ, Arora KK, Mores N, Tomic M, Stojilkovic SS, Catt KJ 2000 Local regulation of gonadotroph function by pituitary gonadotropin-releasing hormone. Endocrinology 141:1187–1195

    Yahalom D, Rahimipour S, Koch Y, Ben-Aroya N, Fridkin M 2000 Design and synthesis of potent hexapeptide and heptapeptide gonadotropin-releasing hormone antagonists by truncation of a decapeptide analogue sequence. J Med Chem 43:2831–2836

    Evans MJ, Kitson NE, Alexander SL, Irvine CH, Turner JE, Perkins NR, Livesey JH 2002 Effectiveness of an antagonist to gonadotrophin releasing hormone on the FSH and LH response to GnRH in perifused equine pituitary cells, and in seasonally acyclic mares. Anim Reprod Sci 73:37–51

    Leung PC, Cheng CK, Zhu XM 2003 Multi-factorial role of GnRH-I and GnRH-II in the human ovary. Mol Cell Endocrinol 202:145–153

    Dekel N, Shalgi R 1987 Fertilization in vitro of rat oocytes undergoing maturation in response to a GnRH analogue. J Reprod Fertil 80:531–535

    Tsafriri A, Adashi E.Y. 1994 Local nonsteroidal regulators of ovarian function. In: Knobil E, Neil JD, eds. The physiology of reproduction, 2nd Ed. New York: Raven Press; 817–860

    Morales P 1998 Gonadotropin-releasing hormone increases ability of the spermatozoa to bind to the human zona pellucida. Biol Reprod 59:426–430

    Corbin A, Bex FJ 1981 Luteinizing hormone releasing hormone agonists induce ovulation in hypophysectomized proestrous rats: direct ovarian effect. Life Sci 29:185–192

    Ekholm C, Hillensjo T, Isaksson O 1981 Gonadotropin releasing hormone agonists stimulate oocyte meiosis and ovulation in hypophysectomized rats. Endocrinology 108:2022–2024

    Bex FJ, Corbin A 1984 Cyclic response of hypophysectomized rats to ovulation induced by LHRH agonist: mediation by prostaglandins. Life Sci 35:969–979

    Hsueh AJ, Jones PB 1981 Extrapituitary actions of gonadotropin-releasing hormone. Endocr Rev 2:437–461

    Mahesh VB, Brann DW 1998 Regulation of the preovulatory gonadotropin surge by endogenous steroids. Steroids 63:616–629

    Koch Y, Baram T 1976 Convenient procedure for extraction of gonadotropin-releasing hormone and thyrotropin-releasing hormone from hypothalamic tissue. FEBS Lett 67:186–188

    Koch Y, Wilchek M, Fridkin M, Chobsieng P, Zor U, Lindner HR 1973 Production and characterization of an antiserum to synthetic gonadotropin-releasing hormone. Biochem Biophys Res Commun 55:616–622

    Koch Y, Chobsieng P, Zor U, Fridkin M, Lindner HR 1973 Suppression of gonadotropin secretion and prevention of ovulation in the rat by antiserum to synthetic gonadotropin-releasing hormone. Biochem Biophys Res Commun 55:623–629

    Vandesompele J, De Preter K, Pattyn F, Poppe B, Van Roy N, De Paepe A, Speleman F 2002 Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol 3:34.1–34.12

    Andersen CL, Jensen JL, Orntoft TF 2004 Normalization of real-time quantitative reverse transcription-PCR data: a model-based variance estimation approach to identify genes suited for normalization, applied to bladder and colon cancer data sets. Cancer Res 64:5245–5250

    Stahlberg A, Hakansson J, Xian X, Semb H, Kubista M 2004 Properties of the reverse transcription reaction in mRNA quantification. Clin Chem 50:509–515

    Szabo A, Perou CM, Karaca M, Perreard L, Quackenbush JF, Bernard PS 2004 Statistical modeling for selecting housekeeper genes. Genome Biol 5:R59.1–R59.10

    Pfaffl MW, Tichopad A, Prgomet C, Neuvians TP 2004 Determination of stable housekeeping genes, differentially regulated target genes and sample integrity: BestKeeper-Excel-based tool using pair-wise correlations. Biotechnol Lett 26:509–515

    Gore AC, Roberts JL 1995 Regulation of gonadotropin-releasing hormone gene expression in the rat during the luteinizing hormone surge. Endocrinology 136:889–896

    Suzuki M, Nishihara M, Takahashi M 1995 Hypothalamic gonadotropin-releasing hormone gene expression during rat estrous cycle. Endocr J 42:789–796

    Porkka-Heiskanen T, Urban JH, Turek FW, Levine JE 1994 Gene expression in a subpopulation of luteinizing hormone-releasing hormone (LHRH) neurons prior to the preovulatory gonadotropin surge. J Neurosci 14:5548–5558

    Park OK, Gugneja S, Mayo KE 1990 Gonadotropin-releasing hormone gene expression during the rat estrous cycle: effects of pentobarbital and ovarian steroids. Endocrinology 127:365–372

    Zoeller RT, Young III WS 1988 Changes in cellular levels of messenger ribonucleic acid encoding gonadotropin-releasing hormone in the anterior hypothalamus of female rats during the estrous cycle. Endocrinology 123:1688–1689

    Araki S, Ferin M, Zimmerman EA, Vande Wiele RL 1975 Ovarian modulation of immunoreactive gonadotropins-releasing hormone (Gn-RH) in the rat brain: evidence for a differential effect on the anterior and mid-hypothalamus. Endocrinology 96:644–650

    Levine JE, Ramirez VD 1982 Luteinizing hormone-releasing hormone release during the rat estrous cycle and after ovariectomy, as estimated with push-pull cannulae. Endocrinology 111:1439–1448

    Sarkar DK, Chiappa SA, Fink G, Sherwood NM 1976 Gonadotropin-releasing hormone surge in pro-oestrous rats. Nature 264:461–463

    Jennes L, McShane T, Brame B, Centers A 1996 Dynamic changes in gonadotropin releasing hormone receptor mRNA content in the mediobasal hypothalamus during the rat estrous cycle. J Neuroendocrinol 8:275–281

    Zmeili SM, Papavasiliou SS, Thorner MO, Evans WS, Marshall JC, Landefeld TD 1986 and luteinizing hormone ? subunit messenger ribonucleic acids during the rat estrous cycle. Endocrinology 119:1867–1869

    Prevot V, Dutoit S, Croix D, Tramu G, Beauvillain JC 1998 Semi-quantitative ultrastructural analysis of the localization and neuropeptide content of gonadotropin releasing hormone nerve terminals in the median eminence throughout the estrous cycle of the rat. Neuroscience 84:177–191

    Ortolano GA, Haisenleder DJ, Dalkin AC, Iliff-Sizemore SA, Landefeld TD, Maurer RA, Marshall JC 1988 Follicle-stimulating hormone ? subunit messenger ribonucleic acid concentrations during the rat estrous cycle. Endocrinology 123:2946–2948

    Bauer-Dantoin AC, Hollenberg AN, Jameson JL 1993 Dynamic regulation of gonadotropin-releasing hormone receptor mRNA levels in the anterior pituitary gland during the rat estrous cycle. Endocrinology 133:1911–1914

    Funabashi T, Brooks PJ, Weesner GD, Pfaff DW 1994 Luteinizing hormone-releasing hormone receptor messenger ribonucleic acid expression in the rat pituitary during lactation and the estrous cycle. J Neuroendocrinol 6:261–266

    Savoy-Moore RT, Schwartz NB, Duncan JA, Marshall JC 1980 Pituitary gonadotropin-releasing hormone receptors during the rat estrous cycle. Science 209:942–944

    Clayton RN, Solano AR, Garcia-Vela A, Dufau ML, Catt KJ 1980 Regulation of pituitary receptors for gonadotropin-releasing hormone during the rat estrous cycle. Endocrinology 107:699–706

    Meidan R, Koch Y 1981 Variations in luteinizing hormone-releasing hormone receptors in pituitary cells from immature and mature cycling female rats. FEBS Lett 132:114–116

    Aiyer MS, Chiappa SA, Fink G 1974 A priming effect of luteinizing hormone releasing factor on the anterior pituitary gland in the female rat. J Endocrinol 62:573–588

    Bauer-Dantoin AC, Jameson JL 1995 Gonadotropin-releasing hormone receptor messenger ribonucleic acid expression in the ovary during the rat estrous cycle. Endocrinology 136:4432–4438

    Wong TT, Zohar Y 2004 Novel expression of gonadotropin subunit genes in oocytes of the gilthead seabream (Sparus aurata). Endocrinology 145:5210–5220

    Roth C, Schricker M, Lakomek M, Strege A, Heiden I, Luft H, Munzel U, Wuttke W, Jarry H 2001 Autoregulation of the gonadotropin-releasing hormone (GnRH) system during puberty: effects of antagonistic versus agonistic GnRH analogs in a female rat model. J Endocrinol 169:361–371(Tamar D. Schirman-Hildesh)