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Endocrinology Vol. 141, No. 5 1618-1626
Copyright © 2000 by The Endocrine Society

ARTICLES

Thyroid Hormone and Estrogen Regulate Brain Region-Specific Messenger Ribonucleic Acids Encoding Three Gonadotropin-Releasing Hormone Genes in Sexually Immature Male Fish, Oreochromis niloticus1

Ishwar S. Parhar, Tomoko Soga and Yasuo Sakuma

Department of Physiology, Nippon Medical School, Tokyo 113-8602, Japan

Address all correspondence and requests for reprints to: Dr. Ishwar S. Parhar, Department of Physiology, Nippon Medical School, Sendagi-1–1-5, Bunkyo-ku, Tokyo 113-8602, Japan. E-mail: ishwar{at}nms.ac.jp


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The present study was undertaken to determine whether T3, estrogen, and 11-ketotestosterone could alter a specific population of GnRH-containing neurons, as indicated by a change in messenger RNA (mRNA) levels in sexually immature male tilapia, Oreochromis niloticus. Two weeks after castration, fish were assigned to four treatment groups. One group served as the control (sesame oil); a single ip injection of (T3; 5 µg/g), estradiol benzoate (EB; 5 µg/g), or 11-ketotestosterone (KT; 5 µg/g) was administered to the remaining three groups. Twenty-four hours after the injection, brains were collected and processed for in situ hybridization histochemistry using 35S-labeled 30-mer antisense oligonucleotide probes complementary to the GnRH-coding region of chicken II, salmon, and seabream GnRH. Computerized image analysis was performed to quantify mRNA concentrations, neuronal numbers, and neuronal size of the terminal nerve-nucleus olfactoretinalis, preoptic, and midbrain GnRH neurons. KT had no effect on any of the above neuronal parameters examined for salmon or seabream GnRH. Neither T3, EB, nor KT was effective to induce changes in midbrain chicken GnRH II mRNA concentrations, neuronal numbers, and neuronal size, indicating that an as yet unknown regulatory mechanism may operate midbrain GnRH neurons. T3 specifically suppressed the concentration of terminal nerve salmon GnRH mRNA, and EB significantly increased preoptic seabream GnRH neuronal numbers. These results are consistent with the hypothesis that thyroid hormone, by suppressing terminal nerve GnRH expression, promotes inhibition of sexual maturation. Furthermore, the failure of KT, a nonaromatizable androgen, to influence preoptic GnRH neurons emphasizes that an estrogenic pathway, at the onset of sexual maturation, is responsible for the recruitment of additional preoptic GnRH neurons that are fundamental to reproduction and behavior.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GnRH IS SYNTHESIZED during embryonic development, well in advance of sexual maturation, as evidenced by the presence of GnRH messenger RNA (mRNA) and protein within the brain at early embryonic ages in mammalian and nonmammalian vertebrate species (1, 2, 3, 4, 5). However, GnRH neurons at the onset of sexual maturation await appropriate chemical cues, which, besides a constellation of signals, include responsiveness to feedback by sex steroids (6, 7, 8, 9, 10, 11, 12, 13). Less is known about the interaction between thyroid hormone and GnRH gene expression in vertebrate species. There is, however, compelling evidence in the sheep that emphasizes the role of thyroid hormone in reproduction that could involve the GnRH neuronal system (14, 15, 16). Furthermore, in rodents, hyperthyroidism during prepubertal development prevents maturation of the ovary, and in adults causes ovarian atrophy, cessation of estrous cycles, and alterations in reproductive behaviors (17, 18). In amphibians and salmonids, at the climax of metamorphosis, which coincides with a peak in thyroid hormone, forebrain GnRH neurons develop elaborate neurite arborization and an increase in terminal nerve GnRH peptide concentration (3, 19, 20, 21). More recently, the finding of thyroid hormone receptor colocalized in GnRH neurons raises the intriguing possibility of a direct influence of thyroid hormone on GnRH gene expression (22). Estrogen and thyroid hormone receptors are a superfamily of ligand-activated nuclear transcription factors. To activate transcription, thyroid hormone receptors and estrogen receptors bind to their own specific response element or compete for the common sequence motif in the estrogen and thyroid hormone response elements (23, 24). It is, therefore, reasonable to predict that the influence of thyroid hormone on GnRH gene expression would parallel that of estrogen. However, although it has been known for years that altered estrogen levels effect GnRH neuronal numbers and GnRH peptide and mRNA concentrations, the available information is conflicting (25, 26, 27). Furthermore, it is unclear whether testosterone or its aromatization to estrogen might be involved in the regulation of GnRH mRNA concentrations and neuronal morphology in male fish (28).

Our objective was to determine whether a specific population of GnRH neurons was altered by hormones in sexually immature male tilapia Oreochromis niloticus, a fish species with chicken II, salmon, and seabream GnRH mRNA-synthesizing neuronal cell bodies distributed in the terminal nerve, preoptic area, and midbrain tegmentum, respectively (28, 29). Using quantitative in situ hybridization histochemistry with oligonucleotide DNA probes complementary to chicken II, salmon, and seabream GnRH mRNA, we examined the influence of T3, estrogen, and 11-ketotestosterone (KT; a nonaromatizable androgen predominant in fish) on GnRH mRNA concentrations, GnRH neuronal numbers, and neuronal sizes in castrated sexually immature male tilapia. In addition, we reconstructed the brain to determine the absolute value of GnRH neuronal numbers. Hence, except for our previous study using juvenile bonyfish (30), the present study would provide the first evidence of the role of thyroid hormone, estrogen, and 11-ketotestosterone in the regulation of mRNA of three molecular variants of GnRH localized within the same brain in sexually immature bonyfish.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Sexually immature adult male tilapia Oreochromis niloticus bought from a commercial farmer were allowed to adjust to our laboratory conditions for several weeks. All fish were kept in freshwater under a natural photoperiod (10 h of light, 14 h of darkness). The water temperature was maintained at 26 ± 1 C. The fish were fed commercial fish chow once daily. At the time of death, the average length of fish was 12.5 ± 0.3 cm, the average weight was about 59.1 ± 4.3 g, and the average gonadosomatic index was 0.05. Fish were identified as sexually immature based on the absence of spermatozoa in the testes.

Experimental design
Surgery. Male tilapias were castrated under tricaine methanosulfate anesthesia (MS222, Sigma, St. Louis, MO). A midventral incision was made through the wall of the body cavity, which arched caudoventrally toward the anterior of the urogenital vent. Care was taken to remove the whole length of both testes with a fine forceps. After castration the muscles were stitched back in position using a catgut suture. Five fish were housed in each community tank (length, 60 cm; width, 27 cm; height, 35 cm) containing 40 liters water.

Hormone treatments. Two weeks after the operation, castrated male fish were given a single ip injection of sesame oil (n = 5; Nakarai Tesque, Tokyo) or estradiol benzoate (EB; n = 5; 5 µg/g BW; Sigma) or L-T3 (n = 5; 5 µg/g BW; Sigma) or KT (n = 5; 5 µg/g BW; gift from Dr. Graham Young, Otago University, New Zealand). Hormones were dissolved in 100% ethanol (5 mg/20 µl), then diluted with sesame oil to their final concentration. The selected doses of hormones produced physiological levels of T3, estrogen, and 11-ketotestosterone similar to those of reproductively active male and female cichlids, the tilapia Oreochromis mossambicus and the Haplochromis burtoni [female T3 (31), male T3 (32), female estrogen (33), male estrogen (34), and male KT (35)]. Twenty-four hours after the injection animals were anesthetized by immersing them in a 0.01% solution of MS222 before sacrifice by decapitation. The brains were removed and processed for in situ hybridization. Blood samples were collected to determine levels of estrogen, T3, and KT by RIA.

RIA of estrogen, T3, and KT
Twenty-four hours after hormone injections, blood samples were obtained between 1400 and 1700 h. About 100 µl blood were collected from each fish into heparinized microsyringes from the caudal peduncle. The blood was stored at 4 C for several hours and centrifuged at 4000 rpm to obtain the plasma. The plasma was kept frozen at -80 C. Plasma estrogen, T3, and KT were measured by RIA (36). The assay sensitivity was 50 pg/ml. The results were statistically analyzed using Student’s t test.

In situ hybridization protocol
The in situ hybridization method used has been described previously (37). Briefly, the brains from all experimental groups were fixed in Bouin’s solution for about 18 h. The tissues were then dehydrated through graded ethanols (n-butanol) and embedded in Paraplast Plus (Oxford Labware, St. Louis, MO). Serial sagittal 10-µm sections were mounted onto Silane (Shinestu Silicon Chemicals, Tokyo, Japan)-coated glass microscope slides, dried on a warm plate at 37–42 C, and kept at room temperature until in situ hybridization was carried out. In situ hybridization was performed using commercially synthesized oligonucleotide sense and antisense probes (Greiner Japan, Tokyo, Japan), 30-mer complementary to mRNAs coding for amino acids 1–10 of each preprohormone, that is, the GnRH-coding region of salmon, seabream, and chicken GnRH II (29, 38, 39). The oligonucleotide probes (12 pmol/µl) were 3'-end labeled with [{alpha}-35S]deoxy-ATP (60 pmol; 1250 Ci/mmol; New England Nuclear, Boston, MA) using terminal transferase (25 U; Roche Molecular Biochemicals, Indianapolis, IN) to a specific activity of about 106 cpm/µl. The end-labeled probes were suspended in Tris/EDTA buffer [0.1 M Tris-HCl (pH 7.6), 10 mM triethylamine, and 1 mM EDTA] and isolated through Sephadex G-25 spin columns. A saturating concentration for each of the three GnRH complementary oligonucleotide probes was determined by applying varying amounts of 35S-labeled probe, ranging from 1–20 x 104 cpm/section. Based on the saturation curve, 5 x 104 cpm (salmon GnRH, chicken GnRH II) or 10 x 104 cpm (seabream GnRH) labeled probe in 35 µl hybridization buffer were applied to each tissue section. Control sections were incubated with excess probe or with hybridization buffer lacking the complementary DNA probes. Sections were hybridized at 37 C for 60 h. Then, they were rinsed twice in 2 x SSC (1 x SSC = 0.15 M NaCl and 15 mM sodium citrate) at room temperature for 10 min, in 0.5 x SSC overnight, and in 0.1 x SSC for 6 h. After the wash, all sections were dehydrated, dipped in Kodak NTB3 emulsion (Eastman Kodak Co., Rochester, NY), and stored, desiccated, at 4 C for 10 days. After development in Kodak D-19 developer and fixer, the sections were counterstained with cresyl violet.

Analysis of GnRH neuronal numbers, neuronal sizes, and mRNA concentrations
The terminal nerve GnRH neurons are closely packed and form a ganglia. To determine the volume of the terminal nerve ganglia, the labeled area in every section along the whole rostral-caudal length of the ganglia was measured, and the sum of all of the labeled areas was obtained and multiplied by the section thickness (10 µm). As the preoptic and midbrain GnRH neurons ranged between 20–30 µm2 in diameter, all labeled cells localized in each 30 µm of tissue along the whole rostral-caudal axis of the brain were analyzed. As each brain section contained labeled cells of varying diameter, we therefore analyzed the percentage of GnRH cells for each of the different cell profile areas. Silver grains per cell (per unit area) or per unit volume (terminal nerve ganglia) were quantitated using an MCID image analyzing system (Imaging Research, Inc., Brook University, St. Catharines, Canada) at x300 magnification. For comparisons, previously reported data for untreated fish from the present experiment (38) were used for statistical analyses with Duncan’s new multiple range test.

The results of image analyses were reexamined by drawing each section containing labeled cells. The outline of all the brain sections and the positions of all preoptic and midbrain GnRH cells were charted on the drawings with the aid of a camera lucida attached to an Olympus Corp. light microscope (New Hyde Park, NY) at x4 magnification. The brains were reconstructed to determine the "true" cell numbers.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Animals treated with T3, EB, or KT had significantly elevated, but physiological, levels of plasma estrogen (EB, 3.6 ± 0.9 ng/ml; oil, 0.4 ± 0.1 ng/ml), T3 (T3, 4.6 ± 1.2 ng/ml; oil, 0.2 ± 0.07 ng/ml), and KT (0.9 ± 0.3 ng/ml; oil, 0.1 ± 0.06 ng/ml) compared with oil-treated fish (P < 0.0001, by Student t test).

Using the probes directed against mRNAs of the three GnRH variants, no coexpression was observed; instead, each probe specifically labeled cells in only one of the three brain regions known to express GnRH. Sections from oil-, EB-, T3-, and KT-treated groups displayed a similar distribution of GnRH cells. Salmon GnRH mRNA-expressing cell bodies were seen as a terminal nerve ganglia (nucleus olfactoretinalis) at the caudalmost part of the olfactory bulbs, seabream GnRH mRNA-containing cells in the preoptic area, and chicken GnRH II mRNA-containing cells in the midbrain tegmentum ( Figs. 1–3GoGoGo). These results are consistent with our earlier observations using in situ hybridization and immunocytochemistry (28, 29, 37).



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  		Figure 1. A schematic sagittal brain section representing the distribution of GnRH mRNA-expressing neurons. NOR, Nucleus olfactoretinalis-synthesizing salmon GnRH mRNA; POA, preoptic area-synthesizing seabream GnRH mRNA; MB, midbrain tegmentum-synthesizing chicken GnRH II mRNA.

 


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  		Figure 2. Darkfield photomicrographs of silver grain clusters over cells in the terminal nerve ganglia after hybridization with 35S-labeled oligonucleotide probe complementary to salmon GnRH in oil (A) and T3-treated animals (B). Note the decrease in hybridization signals after T3 treatment (B). Scale bar, 50 µm.

 


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  		Figure 3. Darkfield photomicrographs of preoptic seabream GnRH mRNA-containing cells after oil (A), EB (B), and KT treatment (C). Preoptic GnRH cells are smaller in size than midbrain GnRH cells (D–F). Midbrain chicken GnRH II mRNA containing cells after oil (D), EB (E), and KT (F) treatments. Hormonal manipulations show no effect on chicken GnRH II mRNA content. Scale bar, 50 µm.

 
Terminal nerve neurons (salmon GnRH mRNA)
The number of silver grains per unit volume (grains per mm3) of oil-treated animals was 11.6 ± 2.3 x 106 grains/mm3, and this was not different from that of EB-treated animals (9.9 ± 1.2 x 106 grains/mm3) or KT-treated animals (11.0 ± 2.7 x 106 grains/mm3). The number of silver grains per unit volume decreased significantly after T3 treatment (4.5 ± 0.7 x 106 grains/mm3; P < 0.01 compared with oil and KT; P < 0.05 compared with EB; Figs. 2Go and 4Go, left panel).



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  		Figure 4. Histograms showing the total volume (right panel) and the cellular salmon GnRH mRNA content (left panel) of the terminal nerve ganglia. NOR, Nucleus olfactoretinalis; OIL, sesame oil. *, P < 0.01 significantly different from oil and KT; P < 0.05 compared with EB (by Duncan’s new multiple range test).

 
The total volume of the terminal nerve neurons (ganglia) containing salmon GnRH mRNA (right and left halves included) was not different between oil (11.6 ± 2.1 x 10-4 mm3), EB (8.9 ± 2.2 x 10-4 mm3), T3 (13.2 ± 0.6 x 10-4 mm3), and KT (9.3 ± 0.4 x 10-4 mm3) treatment groups (Fig. 4Go, right panel).

Preoptic neurons (seabream GnRH mRNA)
In the oil-, EB-, and T3-treated groups the distribution of cells expressing seabream GnRH mRNA was skewed to the left, indicating that majority of cells have lower grain counts, whereas in the KT-treated animals, GnRH neurons were seen expressing different densities of seabream GnRH mRNA, with almost equal numbers of cells having grain densities ranging from 1–45 grains x 104/mm2·cell, as represented in Fig. 5Go (left panel). However, when the mean values of silver grain densities were combined to give experimental group means, there was no change in the total grain counts after EB, T3, or KT treatment (oil, 12.8 ± 2.1; EB, 15.7 ± 1.7; T3, 10.2 ± 2.0; KT, 15.0 ± 5.0 grains x 104/mm2·cell; Figs. 3Go, A–C, and 5, left panel inset).



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  		Figure 5. Histograms displaying the sizes (right panel) and the percentage of labeled cells expressing different levels of seabream GnRH mRNA (left panel). The mean size of preoptic GnRH cells and the mean seabream GnRH mRNA content (±SE) per brain are shown in the box of the right and left panels, respectively. No statistical difference was seen in any of these measures between treatment groups.

 
Each brain section of the preoptic area contained GnRH cells of different sizes ranging from 60–450 µm2 for the oil-treated, 50–380 µm2 for the EB-treated, 60–330 µm2 for the T3-treated, and 50–400 µm2 for the KT-treated group. The percentage of GnRH cell profiles with small surface area increased after EB (median, 160) and T3 (median, 160) treatments, with a distribution slightly skewed to the left of the oil- and KT-treated groups (median, 200; Fig. 5Go, right panel). However, when the mean values of cell profile area was combined to give experimental group means, there was no significant difference in the total cell profile area between experimental groups (oil, 202.8 ± 17.9; EB, 168.6 ± 9.4; T3, 169.2 ± 9.1; KT, 191.6 ± 18.9 µm2; Fig. 5Go, right panel inset). In contrast, the total number of cells expressing seabream GnRH mRNA per brain, compared with those in the oil, T3, and KT treatment groups, increased significantly after EB treatment (oil, 49.3 ± 15.3; EB, 138. 8 ± 23.1; T3, 34.6 ± 18.7; KT, 80.3 ± 44.4 cells; P < 0.01; Fig. 6Go, left panel). Similarly, when the total number of cells expressing seabream GnRH mRNA per brain were counted after reconstruction of tracings of brain sections, there was a significant increase after EB treatment (oil, 21.3 ± 2.4; EB, 54.5 ± 5.5; T3, 12.6 ± 5.4; KT, 48.7 ± 21.2 cells; P < 0.01). In some instances, KT treatment caused extreme variation in GnRH cell numbers and mRNA levels that apparently shifted the distribution to the right and increased the SE, which explains the statistically insignificant influence of KT (Figs. 5Go and 6Go, left panel).



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  		Figure 6. The mean number of midbrain chicken GnRH II (right panel) and preoptic seabream GnRH cells (left panel). Hormonal manipulations had no effect on the mean numbers of midbrain GnRH cells. The total number of preoptic GnRH cells per brain increased significantly after EB treatment. *, P < 0.01 significantly different from oil, T3, and KT treatment (by Duncan’s new multiple range test).

 
Midbrain neurons (chicken GnRH II mRNA)
In the oil-treated animals, GnRH neurons expressed different densities of chicken GnRH II mRNA, with almost equal numbers of cells having grain densities ranging from 1–48 grains x 104/mm2·cell, as represented in Fig. 7Go (left panel). After EB, T3, and KT treatment, the number of cells expressing low cellular GnRH mRNA increased, with a distribution significantly skewed to the left of that in the oil-treated group (Fig. 7Go, left panel). However, when the mean silver grain densities were combined to give experimental group means, there was no change in the total grain counts after EB, T3, or KT treatment (oil, 19.9 ± 4.4; EB, 12.2 ± 1.8; T3, 15.0 ± 2.7; KT, 15.8 ± 1.6 grains x 104/mm2·cell; Fig. 7Go, left panel inset).



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  		Figure 7. Histograms displaying the sizes (right panel) and the percentage of labeled cells expressing different levels of chicken GnRH II mRNA (left panel). The distribution of cells after EB, T3, and KT treatment is skewed to the left of the oil-treated group, indicating more cells with low cellular GnRH mRNA content (left panel). However, no statistical difference was seen in the mean size of midbrain GnRH cells and the mean chicken GnRH II mRNA content (±SE) per brain as shown in the box of the right and left panels, respectively.

 
In all experimental groups, each brain section of the midbrain area contained GnRH cells of different sizes, ranging from 60–920 µm2 (median, 360 µm2; Fig. 7Go, right panel). When the mean cell profile areas were combined to give experimental group means, there was no significant difference in the total cell profile area among experimental groups (oil, 375.3 ± 18.7; EB, 390.9 ± 20.5; T3, 408.8 ± 13.6; KT, 382.2 ± 29.7 µm2; Fig. 7Go, right panel inset). There was no difference in the total number of cells expressing chicken GnRH II mRNA per brain among treatment groups (oil, 70.8 ± 17.6; EB, 68.3 ± 6.1; T3, 71.2 ± 14.8; KT, 80.3 ± 15.2 cells; Fig. 6Go, right panel). Furthermore, the total number of cells expressing chicken GnRH II mRNA per brain, counted after reconstruction of tracings of brain sections, showed no significant difference (oil, 38.3 ± 7.3; EB, 39.0 ± 4.4; T3, 38.6 ± 8.4; KT, 47.5 ± 4.4 cells).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In the present study we examined whether T3, EB, and KT could alter a specific population of GnRH mRNA-containing neurons in the sexually immature male tilapia. The number of neurons expressing GnRH mRNA and the relative degree of expression were analyzed using in situ hybridization and quantitative image analysis with deoxyoligonucleotide probes complementary to the cichlid GnRH-coding region (29, 37, 38, 39). The specificity of our method to detect salmon GnRH mRNA in the terminal nerve-nucleus olfactoretinalis, seabream GnRH mRNA in the preoptic area, and chicken GnRH II mRNA in the midbrain is evidenced by the similarity in distribution of our labeled cells with previous immunocytochemical and in situ hybridization studies as well as the absence of positively labeled cells in sections treated with excess probes or with hybridization buffer lacking the complementary DNA probes (28, 37). Several important conclusions about the regulation of brain region-specific GnRH mRNA-containing neurons by T3, EB, and KT can be drawn from this experiment. First, the most dramatic effect of T3 was observed in the terminal nerve ganglia, in which there was a pronounced decrease in GnRH labeling intensity. Second, although manipulations of EB significantly increased the number of labeled preoptic GnRH cells, KT treatment had the tendency to increase their labeling intensity. Finally, T3, EB, and KT treatments had no influence on the midbrain GnRH mRNA content or neuronal numbers, indicating that another regulatory pathway might operate chicken GnRH II-containing neurons.

Terminal nerve neurons (salmon GnRH mRNA)
In amphibians and salmonids at the climax of metamorphosis, which coincides with a peak in thyroid hormone, forebrain GnRH neurons develop elaborate neurite arborization and GnRH peptide expression, followed by a rise in plasma testosterone and estrogen levels (3, 19, 20, 21). The present study further emphasizes that the terminal nerve GnRH neurons are influenced by T3 treatment, which significantly lowers copies of cellular salmon GnRH mRNA in sexually immature tilapia. Considering these results, it is logical to postulate that the effect of thyroid hormone on the terminal nerve GnRH neurons can have consequences on reproductive functions. In this context, it is worth noting that overexpression (hyperthyroidism) of thyroid hormone during prepubertal development in rodents prevents maturation of the ovary (17) and causes anestrus in the adult sheep (15). Interestingly, in the tilapia, the expression of terminal nerve salmon GnRH first begins on day 10 ± 1 (29), which coincides with a significant decrease in thyroid hormones (40). Therefore, it is reasonable to assume that during early development in the tilapia, the high concentrations of endogenous thyroid hormone (31), independently or in concert with neurotransmitters/neuromodulators ({gamma}-aminobutyric acid, opioids, neuropeptide Y, galanin, and neurotensin) (9, 10), could suppress the onset of sexual maturation by inhibiting the terminal nerve GnRH neurons. On the other hand, the initiation of maturation or reproductive activity is the result of a decrease in thyroid hormone as seen in salmonids (3, 20, 41) and sheep (14). Interestingly, sexually mature tilapia have low levels of thyroid hormones (31, 32, 40) but high levels of terminal nerve salmon GnRH mRNA (28) relative to tilapia treated with thyroid hormone (present study). What remains uncertain is the mechanism by which thyroid hormone regulates the inhibition of salmon GnRH mRNA during sexually immature stages. Several assumptions can be considered. First, it is possible that T3 activates GnRH neurons or GnRH target tissues involved in the maturation of the hypothalamo-hypophyseal-gonadal axis in sexually immature animals and thereby increases the demand for GnRH peptide. Therefore, the decrease in salmon GnRH message could be the result of rapid translation of message to GnRH peptide synthesis/secretion. However, it is also possible that thyroid hormone could act directly on terminal nerve GnRH neurons to decrease their synthetic capacity or promote degradation of GnRH mRNA, but it remains to be seen whether terminal nerve GnRH neurons possess thyroid hormone response element and/or thyroid hormone receptors to qualify as an alternative pathway. Thyroid hormone could act via estrogen response element, reported in the promoter of the terminal nerve salmon GnRH gene in the salmonids (Onchorynchus nerka) (42). Apparently, estrogen response element is either absent in the tilapia salmon GnRH gene or it is nonoperational during sexually immature stages, as EB administration had no effect on salmon GnRH mRNA. A more likely scenario would be that the expression of terminal nerve GnRH is mediated by an intermediate agent. There is evidence in mammals suggesting that thyroid hormones and reproductive hormones are functionally linked insofar as thyroid hormone stimulates testosterone production (43) and the onset of puberty (44). Further, our previous study showed that T3 stimulates testosterone production in the tilapia (34) and that testosterone (28), but not KT (nonaromatizable androgen) (45) or estrogen (unpublished observations), lead to an increase in terminal nerve salmon GnRH mRNA in sexually mature male tilapia. Therefore, based on these findings we postulate that T3 by regulating androgen receptors (46) could influence the androgen responsiveness of the terminal nerve GnRH neurons during sexually immature stages (Fig. 8Go).



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  		Figure 8. A schematic representation of GnRH neurons in the terminal nerve-nucleus olfactoretinalis (NOR) and the preoptic area (POA). Thyroid hormone (T3) and testosterone (T) could influence the NOR GnRH gene expression through thyroid hormone receptors (THR) and androgen receptors (AR). The POA GnRH gene expression is regulated by estrogen and probably by NOR neuronal fibers morphologically seen traversing the preoptic area.

 
Preoptic neurons (seabream GnRH mRNA)
Estrogen has been demonstrated to regulate transcription of the GnRH gene, but reports concerning the concentrations of preoptic GnRH peptide and mRNA, and GnRH neuronal numbers are conflicting (25, 26, 47). In the present study, considering that replacement with KT, a nonaromatizable androgen, had no effect on preoptic GnRH neuronal cytology or total GnRH mRNA levels supports the inhibitory effect of testosterone on preoptic seabream GnRH mRNA in the tilapia (28), and the androgen-induced increase in neuronal size and neuronal numbers in bonyfish (35, 48) is probably mediated through an aromatase pathway. Furthermore, replacement with EB, but not KT, dramatically increased the total number of detectable preoptic GnRH-expressing neurons, which equaled those in sexually mature tilapia (28), suggesting that estrogen-induced positive feedback in sexually immature fish, might involve recruitment of additional, previously low (or non-) expressing GnRH neurons as reported in rodents (49, 50, 51). However, we and others (50) did not observe a marked increase in detectable levels of preoptic GnRH mRNA, much as has been reported in some mammalian species (25, 26, 47). Our inability to observe an estrogen-induced significant increase in preoptic GnRH mRNA may reflect one of two possibilities. First, GnRH is induced, but to a lower degree of activation. Second, we may have missed the peak of GnRH mRNA expression, at a sampling time not examined in this study. In any case, our data do support the assumption that the neurons of the preoptic area are the population of GnRH neurons that are responsive to estrogen. The pathways through which estrogen influences GnRH gene transcription are not yet established in vertebrates. There is no positive evidence for the expression of classical estrogen receptors in GnRH neurons of bonyfish (52). The lack of estrogen receptor immunoreactivity alone is, however, not definitive evidence for the absence of estrogen receptors in GnRH neurons, as shown recently in rodents; this could be due to technical difficulties (53). We, therefore, do not preclude a direct effect of estrogen on preoptic neurons because of the presence of estrogen response elements in the promoter of preoptic seabream GnRH gene (54). Also, it remains possible that preoptic GnRH neurons express estrogen receptors during early embryogenesis and that estrogen receptor synthesis is abruptly down-regulated and therefore become undetectable because of their low abundance or rapid turnover, as sexual maturation progresses. Furthermore, estrogen receptor subtypes, such as estrogen receptor ß (55, 56) or alternatively spliced forms, might exist in preoptic GnRH neurons of bonyfish that could mediate estrogenic influence. Besides a direct effect, it is possible that estrogen receptor-containing neurons in other brain sites act as intermediaries that could regulate GnRH mRNA via afferent projections to the preoptic GnRH neurons. Neuronal afferents of diverse chemical phenotypes synapsing on GnRH neuronal cell bodies have been implicated in stimulating or inhibiting GnRH mRNA expression in response to estrogen (9, 10, 27, 57). Unfortunately, in bonyfish the limited data demonstrating the afferent projections or colocalization of neuropeptides, androgen and estrogen receptor subtypes, and GnRH do not allow us to distinguish among the above possible interpretations.

Midbrain neurons (chicken GnRH II mRNA)
Our previous study using juvenile and adult castrated male tilapia showed no effect of steroids (T3, estrogen, testosterone, progesterone) on chicken GnRH II cell numbers, cell sizes, and mRNA levels (28, 30, 38). Similarly, in chickens, testosterone has no effect on chicken GnRH II peptide content (58). Therefore, in the tilapia, chicken GnRH II mRNA-synthesizing neurons may not be targets for the feedback effects of steroid hormones. However, it is tempting to speculate that the decrease in chicken GnRH II mRNA, although not significant, might be an estrogenic effect, because androgens in combination with estradiol could decrease chicken GnRH II peptide in the European eel (59). Further, evidence from studies in the musk shrew indicate that chicken GnRH II peptide is modified by ovulation, and that ovarian hormones can modulate some aspects of its production and/or release (60, 61). Although no known function has been described for the chicken GnRH II in teleosts, the chicken GnRH II-containing midbrain neuronal group in mammals (60, 61, 62) has been implicated in reproductive behavior (63). Certainly, further studies are required to test and clarify the regulatory mechanisms of chicken GnRH II. Understanding the expression pattern of the chicken GnRH II gene will be fundamental to elucidate its function in vertebrates.

The present study provides evidence that midbrain chicken GnRH II-containing neurons are insensitive to steroid hormones in sexually immature animals. The study also shows that thyroid hormone promotes inhibition of terminal nerve salmon GnRH expression during sexually immature stages, and the rise in estrogen, at the onset of sexual maturation, recruits additional preoptic seabream GnRH-containing neurons that are fundamental to reproduction. The significance of brain region-specific changes in GnRH mRNA expression in relation to sexual maturation warrants further investigation.


   Acknowledgments
 
We thank Prof. I. Wakabayashi (Nippon Medical School, Tokyo, Japan) for use of the image-analyzing system. We are also grateful to Profs. K. Yamauchi and S. Adachi (Faculty of Fisheries, Hokkaido University, Hakodate, Japan) for their help with the hormone assays.


   Footnotes
 
1 This work was supported by research grants from Nippon Medical School and the Japanese Ministry of Education, Science, Sports, and Culture (Grant 09680801; to I.S.P.). Back

Received December 20, 1999.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Schwanzel-Fukuda M, Pfaff DW 1989 Origin of luteinizing hormone-releasing hormone neurons. Nature 338:161–164[CrossRef][Medline]
  2. Wray S, Grant P, Gainer H 1989 Evidence that cells expressing luteinizing hormone releasing hormone mRNA in the mouse are derived from progenitor cells in the olfactory placode. Proc Natl Acad Sci USA 86:8132–8136[Abstract/Free Full Text]
  3. Parhar IS, Iwata M, Pfaff DW, Schwanzel-Fukuda M 1995 Embryonic development of gonadotropin-releasing hormone neurons in the sockeye salmon. J Comp Neurol 362:256–270[CrossRef][Medline]
  4. Muske LE 1997 Ontogeny, phylogeny and neuroanatomical organization of multiple molecular forms of GnRH. In: Parhar IS, Sakuma Y (eds) GnRH Neurons: Gene to Behavior. Brain Shuppan, Tokyo, pp 145–180
  5. Mulrenin EM, Witkin JW, Silverman A-J 1999 Embryonic development of the gonadotropin-releasing hormone (GnRH) system in the chick: a spatio-temporal analysis of GnRH neuronal generation, site of origin, and migration. Endocrinology 140:422–433[Abstract/Free Full Text]
  6. Ojeda SR, Urbanski HF 1994 Puberty in the rat. In: Knobil E, Neill JD (eds) Physiology of Reproduction. Raven Press, New York, pp 363–409
  7. Foster DL 1994 Puberty in sheep. In: Knobil E, Neill JD (eds) Physiology of Reproduction. Raven Press, New York, pp 411–451
  8. Plant TM 1994 Puberty in primates. In: Knobil E, Neill JD (eds) Physiology of Reproduction. Raven Press, New York, pp 453–485
  9. Kalra SP, Horvath T, Naftolin F, Xu B, Pu S, Kalra PS 1997 The interactive language of the hypothalamus for the gonadotropin-releasing hormone (GnRH) system. J Neuroendocrinol 9:569–576[CrossRef][Medline]
  10. Terasawa E, Nyberg CL 1997 LHRH pulse generation in the monkey: in vivo and in vitro studies. In: Maeda K, Tsukamura H,Yokoyama A (eds) Neural Control of Reproduction, Physiology and Behavior. Karger, Tokyo, pp 57–70
  11. Tang YP, Kashon LM, Sisk CL 1997 Brain region-specific regulation of luteinizing hormone-releasing hormone messenger ribonucleic acid in the male ferret: interactions between pubertal maturation and testosterone. Endocrinology 138:4740–4747[Abstract/Free Full Text]
  12. Martini L, Motta M, Piva F, Zanisi M 1997 LHRF, LHRH, GnRH: what controls the secretion of this hormone? Mol Psychiatry 2:373–376[CrossRef][Medline]
  13. Foster DL, Nagatani S 1999 Physiological perspectives on leptin as a regulator of reproduction: role in timing puberty. Biol Reprod 60:205–215[Abstract/Free Full Text]
  14. Webster JR, Moenter SM, Barrell GK, Lehman MN, Karsch FJ 1991 Role of the thyroid gland in seasonal reproduction. III. Thyroidectomy blocks seasonal suppression of gonadotropin-releasing hormone secretion in sheep. Endocrinology 129:1635–1643[Abstract]
  15. Thrun LA, Dahl GE, Evans NP, Karsch FJ 1997 A critical period for thyroid hormone action on seasonal changes in reproductive neuroendocrine function in the ewe. Endocrinology 138:3402–3409[Abstract/Free Full Text]
  16. Viguie C, Battaglia DF, Krasa HB, Thrun LA, Karsch FJ 1999 Thyroid hormones act primarily within the brain to promote the seasonal inhibition of luteinizing hormone secretion in the ewe. Endocrinology 140:1111–1117[Abstract/Free Full Text]
  17. Young WC 1961 The hormones and mating behavior. In: Young WC (eds) Sex and Internal Secretions. Williams & Wilkins, Baltimore, pp 1210–1212
  18. Dellovade TL, Zhu Y-S, Krey L, Pfaff DW 1996 Thyroid hormone and estrogen interact to regulate behavior. Proc Natl Acad Sci USA 93:12581–12586[Abstract/Free Full Text]
  19. Parhar IS, Koibuchi N, Sakai M, Iwata M, Yamaoka S 1994 Gonadotropin-releasing hormone (GnRH): expression during salmon migration. Neurosci Lett 172:15–18[CrossRef][Medline]
  20. Parhar IS, Iwata M 1996 Intracerebral expression of gonadotropin-releasing hormone and growth hormone-releasing hormone is delayed until smoltification in the salmon. Neurosci Res 26:299–308[CrossRef][Medline]
  21. Hayes WP, Wray S, Battey JF 1994 The frog gonadotropin-releasing hormone-I (GnRH-I) gene has a mammalian-like expression pattern and conserved domains in GnRH-associated peptide, but brain onset is delayed until metamorphosis. Endocrinology 134:1835–1845[Abstract]
  22. Jansen HT, Lubbers LS, Macchia E, Degroot LJ, Lehman MN 1997 Thyroid hormone receptor (a) distribution in hamster and sheep brain: colocalization in gonadotropin-releasing hormone and other identified neurons. Endocrinology 138:5039–5047[Abstract/Free Full Text]
  23. Glass CK, Holloway JM, Devary OV, Rosenfeld MG 1988 The thyroid hormone receptor binder with opposite transcriptional effects to a common sequence motif in thyroid hormone and estrogen response elements. Cell 54:313–323[CrossRef][Medline]
  24. Zhu Y-S, Yen PM, Chin WW, Pfaff DW 1996 Estrogen and thyroid hormone interaction on regulation of gene expression. Proc Natl Acad Sci USA 93:12587–12592[Abstract/Free Full Text]
  25. Sagrillo CA, Grattan DR, McCarthy MM, Selmanoff M 1996 Hormonal and neurotransmitter regulation of GnRH gene expression and related reproductive behaviors. Behav Genet 26:241–277[CrossRef][Medline]
  26. Gore AC, Roberts JL 1997 Regulation of gonadotropin-releasing hormone gene expression in vivo and in vitro. Front Neuroendocrinol 18:209–245[CrossRef][Medline]
  27. Herbison AE 1998 Multimodal influence of estrogen upon gonadotropin-releasing hormone neurons. Endocr Rev 19:302–330[Abstract/Free Full Text]
  28. Soga T, Sakuma Y, Parhar IS 1998 Testosterone differentially regulates expression of GnRH messenger RNAs in the terminal nerve, preoptic and midbrain of male tilapia. Mol Brain Res 60:13–20[Medline]
  29. Parhar IS 1997 GnRH in tilapia: three genes, three origins and their roles. In: Parhar IS, Sakuma Y (eds) GnRH Neurons: Gene to Behavior. Brain Shuppan, Tokyo, pp 99–122
  30. Parhar IS, Soga T, Sakuma Y 1996 In situ hybridization for two differentially expressed GnRH genes following estrogen and triiodothyronine treatment in the brains of juvenile tilapia (cichlid). Neurosci Lett 218:135–138[CrossRef][Medline]
  31. Weber GM, Okimoto DK, Richman III NH, Grau EG 1992 Patterns of thyroxine and triiodothyronine in serum and follicle-bound oocytes of the tilapia, Oreochromis mossambicus, during oogenesis. Gen Comp Endocrinol 85:392–404[CrossRef][Medline]
  32. Sukumar P, Munro AD, Mok EYM, Subburaju S, Lam TJ 1997 Hypothalamic regulation of the pituitary-thyroid axis in the tilapia Oreochromis mossambicus. Gen Comp Endocrinol 106:73–84[CrossRef][Medline]
  33. Yaron Z, Terkatin-Shimony A, Shaham Y, Salzer H 1977 Occurrence and biological activity of estradiol 17-ß in the intact and ovariectomized tilapia aurea (Cichlidae, teleostei). Gen Comp Endocrinol 33:45–52[CrossRef][Medline]
  34. Singh I 1990 Neuroendocrinology of Oreochromis mossambicus. PhD Thesis, Natinoal University of Singapore, Singapore, vol 1:1–265
  35. Soma KK, Francis RC, Wingfield JC, Fernald RD 1996 Androgen regulation of hypothalamic neurons containing gonadotropin-releasing hormone in a cichlid fish: integration with social cues. Horm Behav 30:216–226[CrossRef][Medline]
  36. Young G, Crim LW, Kagawa H, Kambegawa A, Nagahama Y 1983 Plasma 17{alpha},20ß-dihydroxy-4-pregnen-3-one levels during sexual maturation of amago salmon (Oncorhynchus rhodurus): correlation with plasma gonadotropin and in vitro production by ovarian follicles. Gen Comp Endocrinol 51:96–105[CrossRef][Medline]
  37. Parhar IS, Pfaff DW, Schwanzel-Fukuda M 1996 Gonadotropin-releasing hormone gene expression in teleosts. Mol Brain Res 41:216–227[Medline]
  38. Parhar IS, Soga T, Sakuma Y 1998 Quantitative in situ hybridization of three gonadotropin-releasing hormone-encoding mRNAs in castrated and progestrone-treated male tilapia. Gen Comp Endocrinol 112:406–414[CrossRef][Medline]
  39. White SA, Kasten TL, Bond CT, Adelman JP, Fernald RD 1995 Three gonadotropin-releasing hormone genes in one organism suggest novel roles for an ancient peptide. Proc Natl Acad Sci USA 92:8363–8367[Abstract/Free Full Text]
  40. Okimoto DK, Weber GM, Grau EG 1993 The effects of thyroxine and propylthiouracil treatment on changes in body form associated with a possible developmental thyroxine surge during post-hatching development of tilapia, Oreochromis mossambicus. Zool Sci 10:803–811
  41. Tagawa M, Ogasawara T, Sakamoto T, Miura T, Yamauchi K, Hirano T 1994 Thyroid hormone concentrations in the gonads of wild chum salmon during maturation. Fish Physiol Biochem 13:233–240
  42. Klungland H, Anderson O, Kisen G, Alestrom P, Tora L 1993 Estrogen receptor binds to the salmon GnRH gene in a region with long palindromic sequences. Mol Cell Endocrinol 95:147–154[CrossRef][Medline]
  43. Chiao YC, Lee HY, Wang SW, Hwang JJ, Chien CH, Huang SW, Lu CC, Chen, JJ, Tsai SC, Wang PS 1999 Regulation of thyroid hormones on the production of testosterone in rats. J Cell Biochem 73:554–562[CrossRef][Medline]
  44. Bakke JL, Gellert RJ, Lawrence NL 1970 The persistent effects of perinatal hypothyroidism on pituitary, thyroidal and gonadal functions. J Lab Clin Med 76:25–33[Medline]
  45. Soga T, Sakuma Y, Parhar IS 1998 Regulation of molecular variants of GnRH mRNAs by 11-ketotestosterone in the male tilapia. 4th International Congress of Neuroendocrinology. Folia Endocrinol Jpn 74:473 (Abstract P211)
  46. Panno ML, Sisci D, Salerno M, Lanzino M, Pezzi V, Morrone EG, Mauro L, Palmero S, Fugassa E, Ando S 1996 Thyroid hormone modulates androgen and estrogen receptor content in the Sertoli cells of peripubertal rats. J Endocrinol 148:43–50[Abstract]
  47. Kim K, Seong JY, Kang SS, Sun W 1997 Neuroendocrine regulation of GnRH gene expression. In: Parhar IS, Sakuma Y (eds) GnRH Neurons: Gene to Behavior. Brain Shuppan, Tokyo, pp 401–427
  48. Grober MS, Jackson IMD, Bass A 1991 Gonadal steroids affect LHRH preoptic cell number in sex/role changing fish. J Neurobiol 22:734–741[CrossRef][Medline]
  49. 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[Abstract]
  50. Weesner GD, Krey LC, Pfaff DW 1993 {alpha}1 Adrenergic regulation of estrogenic-induced increases in luteinizing hormone-releasing hormone mRNA levels and release. Mol Brain Res 17:77–82[Medline]
  51. Rubin BS, King JC 1994 The number and distribution of detectable luteinizing hormone (LH)-releasing hormone cell bodies changes in association with the preovulatory LH surge in the brain of young but not middle-aged female rats. Endocrinology 134:467–474[Abstract]
  52. Navas JM, Anglade I, Bailhache T, Pakdel F, Breton B, Jeko P, Kah O 1995 Do gonadotrophin-releasing hormone neurons express estrogen receptors in the rainbow trout? A double immunohistochemical study. J Comp Neurol 363:461–474[CrossRef][Medline]
  53. Butler JA, Sjoberg M, Coen CW 1999 Evidence for oestrogen receptor {alpha}-immunoreactivity in gonadotrophin-releasing hormone-expressing neurones. J Neuroendocrinol 11:331–335[CrossRef][Medline]
  54. Chow MM, Kight KE, Gothilf Y, Alok D, Stubblefield J, Zohar Y 1998 Multiple GnRHs present in a teleost species are encoded by separate genes: analysis of the sbGnRH and cGnRH-II genes from the striped bass, Morone saxatilis. J Mol Endocrinol 21:277–289[Abstract]
  55. Das SK, Taylor JA, Korach KS, Paria BC, Dey SK, Lubahn DB 1997 Estrogenic responses in estrogen receptor-a deficient mice reveal a distinct estrogen signaling pathway. Proc Natl Acad Sci USA 94:12786–12791[Abstract/Free Full Text]
  56. Shughrue PJ, Lane MV, Merchenthaler I 1997 Comparative distribution of estrogen receptor-{alpha} and -ß mRNA in the rat central nervous system. J Comp Neurol 388:507–525[CrossRef][Medline]
  57. Silverman A-J, Livne I, Witkin JW 1994 The gonadotropin-releasing hormone (GnRH) neuronal systems: immunocytochemistry and in situ hybridization. In: Knobil E, Neill JD (eds) Physiology of Reproduction. Raven Press, New York, pp 1683–1709
  58. Wilson SC, Gladwell RT, Cunningham FJ 1990 Differential responses of hypothalamic LHRH-I and LHRH-II to castration and gonadal steroid or tamoxifen treatment in cockerels. J Endocrinol 125:139–146[Abstract]
  59. Montero M, Le Belle N, King JA, Millar RP, Dufour S 1995 Differential regulation of the two forms of gonadotropin-releasing hormone (mGnRH and cGnRH-II) by sex steroids in the European female silver eel (Anguilla anguilla). Neuroendocrinology 61:525–535[Medline]
  60. Rissman EF, Li X 1998 Sex differences in mammalian and chicken-II gonadotropin-releasing hormone immunoreactivity in musk shrew brain. Gen Comp Endocrinol 112:346–355[CrossRef][Medline]
  61. Dellovade TL, Ottinger MA, Rissman EF 1995 Mating alters gonadotropin-releasing hormone cell number and content. Endocrinology 136:1648–1657[Abstract]
  62. Kasten TL, White SA, Norton TT, Bond CT, Adelman JP, Fernald RD 1996 Characterization of two new preproGnRH mRNAs in the tree shrew: first direct evidence for mesencephalic GnRH gene expression in a placental mammal. Gen Comp Endocrinol 104:7–19[CrossRef][Medline]
  63. Sakuma Y, Pfaff DW 1980 LHRH in the mesencephalic central grey can potentiate lordosis reflex of female rats. Nature 283:566–567[CrossRef][Medline]



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