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Endocrinology Vol. 145, No. 6 2959-2967
Copyright © 2004 by The Endocrine Society

Evidence That Dynorphin Plays a Major Role in Mediating Progesterone Negative Feedback on Gonadotropin-Releasing Hormone Neurons in Sheep

Robert L. Goodman, Lique M. Coolen, Greg M. Anderson, Steven L. Hardy, Miro Valent, John M. Connors, Maureen E. Fitzgerald and Michael N. Lehman

Department of Physiology and Pharmacology (R.L.G., G.M.A., S.L.H., M.V., J.M.C.), West Virginia University Health Sciences Center, Morgantown, West Virginia 26506-9229; and Department of Cell Biology, Neurobiology, and Anatomy (L.M.C., M.E.F., M.N.L.), University of Cincinnati College of Medicine, Cincinnati, Ohio 45267-0521

Address all correspondence and requests for reprints to: Dr. Robert L. Goodman, Department of Physiology and Pharmacology, Health Sciences Center North, Medical Center Drive, Morgantown, West Virginia 26506-9229. E-mail: bgoodman{at}hsc.wvu.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Endogenous opioid peptides (EOP) mediate progesterone-negative feedback in many species, but the specific EOP systems involved remain unresolved. We first addressed this question in sheep by determining the role of different EOP receptor subtypes in the medial basal hypothalamus (MBH) and preoptic area (POA). Local administration of EOP receptor antagonists to luteal phase ewes indicated that {kappa}-, but not µ- or {delta}-, receptors mediate the inhibition of LH secretion in the MBH. In contrast, both {kappa}- and µ-, but not {delta}-receptor, antagonists increased LH pulse frequency when placed in the POA. We next examined close appositions between dynorphin ({kappa} ligand) and ß-endorphin (µ ligand) containing varicosities and GnRH perikarya in luteal phase ewes using dual immunocytochemistry and light microscopy. Approximately 90% of MBH GnRH neurons had close associations by dynorphin-containing varicosities, but only 40–50% of GnRH perikarya elsewhere had such close associations. In contrast, the percentage of ß-endorphinergic varicosities close to GnRH neurons was similar among all regions. Electron microscopic analysis demonstrated both dynorphinergic synapses and ß-endorphinergic synapses onto GnRH perikarya. These and other data lead to the hypothesis that dynorphin neurons play a major role in progesterone-negative feedback in the ewe and that this inhibition may be exerted directly on GnRH perikarya within the MBH, whereas dynorphin and ß-endorphin input to GnRH neurons in the POA provide redundancy to this system or are involved in other actions of progesterone or estradiol in the control of the GnRH surge.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GNRH NEURONS PLAY a central role in the complex feedback relationships within the hypothalamohypophyseal-ovarian axis that ensure normal ovarian function (1, 2). Two distinct modes of GnRH secretion occur during the ovarian cycle: 1) the preovulatory GnRH surge that drives the LH surge responsible for ovulation and 2) basal, or tonic, GnRH secretion that occurs episodically throughout follicular and luteal phases. The preovulatory GnRH surge is induced by elevated estradiol (1, 2), whereas tonic GnRH secretion is controlled by the negative feedback actions of estradiol and progesterone (1, 2, 3). Specifically, progesterone inhibits GnRH pulse frequency (2, 3, 4, 5) so that slow-frequency pulses are observed during the luteal phase (2, 6, 7), whereas estradiol inhibits GnRH pulse amplitude (8, 9) so that high-frequency, low-amplitude pulses occur in the follicular phase (2, 6, 7).

It is now generally accepted that the inhibitory actions of endogenous opioid peptides (EOPs) play an important role in the control of GnRH secretion. Removal of an EOP brake may play a role in the mechanisms timing the GnRH/LH surge in humans (10), rats (11, 12), and sheep (13). In addition, there is strong evidence in several species (2, 4, 14) that EOPs mediate the negative feedback actions of progesterone. For example, EOP antagonists increase LH (15, 16) and GnRH (17) pulse frequency in luteal-phase and progesterone-treated ovariectomized (OVX) ewes but not in OVX or estradiol-treated OVX females (9, 16). There is also some evidence that EOPs may mediate estradiol-negative feedback on GnRH pulse amplitude (16), but direct measurements of GnRH pulses in OVX and estradiol-treated OVX ewes did not support this hypothesis (9).

Whereas inhibitory effects of EOPs on GnRH pulse frequency and the preovulatory GnRH surge are well established, the role of specific EOP systems is much less clear. Based on the effects of the endogenous ligands (18), receptor-specific agonists (19) and antagonists (20, 21) and ligand immunoneutralization (22, 23), ß-endorphin, and dynorphin acting via µ- and {kappa}-receptors, respectively, are potent physiological inhibitors of LH and GnRH secretion, whereas {delta}receptors appear to play little role in the control of these systems. The ß-endorphin/µ-receptor system appears to be important for control of the GnRH surge (12, 13, 24), but the system mediating progesterone-negative feedback remains controversial (14, 20, 21).

In most mammals, GnRH perikarya are scattered along a rostral-caudal continuum extending from the diagonal Band of Broca, through the preoptic area (POA), to the medial basal hypothalamus (MBH) (25). There is some anatomical evidence for different subpopulations of GnRH neurons (25), and we recently reported that systemic administration of a nonselective EOP antagonist to luteal-phase ewes activated GnRH perikarya located in the MBH but not those in the POA (26). These data raise the possibility that there may be differential EOP regulation of GnRH neurons in the ovine POA and MBH. In this study, we tested this hypothesis by determining whether different subtypes of EOP receptors in the POA and MBH hold LH pulse frequency in check during the luteal phase and analyzing ß-endorphin and dynorphin input to GnRH perikarya in these two regions.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Mature, black-faced ewes of mixed breeding showing regular estrous cycles were maintained in an open barn that allowed exposure to ambient photoperiod and temperature, with access to water and a daily silage allowance. All experiments were done during the breeding season (from late September through December). Surgeries and perfusions were performed in an indoor animal facility. Ewes were moved 3–7 d before the procedures and housed two per pen under artificial lighting that was similar to that outdoors. Ewes were kept in this facility for approximately 10–14 d after surgery so that their recovery could be closely monitored and then returned to the open-sided barn for experimental procedures. Blood samples were collected by jugular venipuncture, allowed to clot over night, and serum harvested and stored at –20 C until assayed. All procedures involving animals were approved by the West Virginia University Animal Care and Use Committee.

Surgery
We stereotaxically implanted bilateral 18-gauge stainless steel guide tubes aimed at either the MBH (n = 8) or POA (n = 11) in experiments 1 and 2, respectively, using sterile techniques, with ewes under gas anesthesia (4 oxygen/1 nitrous oxide, supplemented with halothane at 1–4%) as previously described (27, 28). Briefly, a 2-cm diameter hole was drilled in the skull, the sagittal sinus ligated, and radioopaque dye injected into one lateral ventricle. Positive ventrilography in the sagittal plane (experiment 1) and both sagittal and coronal planes (experiment 2) was used to place the tips of the guide tubes 2 mm dorsal to the target area and 1–1.5 mm (MBH) or 2–2.5 mm (POA) lateral to midline. After protecting the surface of the brain with gel foam and a wire mesh, guide tubes were cemented in place with cranial screws and dental acrylic, plugged with 22-gauge wire stylets, and protected with a plastic cap.

Experimental protocols
We determined which EOP receptor subtype actively inhibited LH pulse frequency during the luteal phase by local administration of antagonists to the {delta} (naltrindole), {kappa} (nor-binaltorphimine [nor-BNI]) and µ (naloxonazine) EOP receptors (29) purchased from Research Biochemical International (Natick, MA); naloxone (Sigma Chemical Co., St. Louis, MO) was used as a positive control. Naloxone is a relative nonspecific EOP receptor antagonist [IC50 for {delta}: 68 nM, {kappa}: 0.7 nM, µ: 1.7 nM (30)] that acts in both the POA and MBH to stimulate LH secretion in luteal-phase ewes (23).

Ovarian cycles of all ewes were synchronized by im injections of 10 mg prostaglandin F-2{alpha} (Luteolyse, Pharmacia & Upjohn, Kalamazoo, MI) spaced 1 wk apart (31) and experimental treatments begun 8 d after the second injection (d 6 of estrous cycle). The day before treatment, crystalline antagonist was tamped into the lumen of sterile 22-gauge tubing cut to extend 2 mm beyond the end of the guide tubes. The next day blood samples were collected every 12 min for 36 min before and 4 h after insertion of antagonist-filled, or empty (negative control), tubing through the guide tubes into either the MBH or POA. Microimplants were then removed, stylets replaced, and two 5-mg prostaglandin F-2{alpha} injections given im 3 h apart to induce luteolysis. This treatment protocol was then repeated four more times, with 8 d between each treatment so that all animals received all treatments; order of treatments was determined using a Latin Square design and did not affect the response to specific antagonists (data not shown). At the end of the experiments, animals were euthanized with an iv injection of approximately 2 g sodium pentobarbital (Sigma) in 0.1 M phosphate buffer (PB) and tissue collected for histological determination of implantation sites (see below for details).

Tissue collection
For light microscopic analyses of tissue (experiments 1–3), ewes were injected twice before perfusion (10 min apart) with 25,000 U heparin iv, followed by sodium pentobarbital (2 g iv). When the animal had stopped breathing, we quickly removed its head and perfused the brain via both internal carotid arteries with 6 liters of 4% paraformaldehyde in 0.1 M PB containing 10 U heparin per milliliter and 0.1% sodium nitrite. A block of tissue containing preoptic and hypothalamic tissue was dissected out, incubated in the same fixative overnight (4 C), and then infiltrated with 30% sucrose in PB. After the tissue had sunk (indicating sucrose infiltration), thick coronal sections (55 µm) were cut using a freezing microtome. For animals that had been treated with receptor antagonists (experiments 1 and 2), a series of every fourth section was mounted on slides, stained with cresyl violet, and examined to determine implant location. For analysis of EOP varicosities close to GnRH neurons, tissue from three additional untreated luteal-phase ewes was collected, sectioned, and four separate series (220 µm apart) stored at –20 C in cryopreservative (32) until processed for dual immunocytochemistry (ICC).

For electron microscopic (EM) analyses (experiment 4), a separate group of luteal-phase ewes (n = 3) were anesthetized (approximately 1 g pentobarbital iv) and intracranially perfused via both carotid arteries with 4% paraformaldehyde containing 0.2% glutaraldehyde and heparin as previously described (33). The preoptic-hypothalamic tissue block was dissected out and incubated in fixative at 4 C overnight and then in stored PB at 4 C. Coronal sections (50 µm) through both the POA and the MBH were cut on a vibratome and stored at –20 C in a cryopreservative solution until processed for dual-label EM immunocytochemistry.

Analytical procedures
LH concentrations were measured in duplicate 200-µl aliquots of serum by RIA as previously described (16); assay sensitivity averaged 0.4 ng/ml (National Institutes of Health S24) and inter- and intraassay coefficients of variation of a pool that produced 60% displacement of iodinated LH were 8.5 and 10.1%, respectively. Progesterone was measured in selected samples to confirm ewes were in the luteal phase using a commercially available kit (28); assay sensitivity averaged 0.03 ng/ml.

We used two separate series of coronal sections, 220 µm apart, to examine the number of dynorphin and ß-endorphin containing varicosities close to GnRH perikarya at the light level. Free-floating tissue sections were processed for dual ICC as previously described (34). Briefly, dynorphin or ß-endorphin was first detected using a modified avidin-biotin-immunoperoxidase protocol in which nickel-enhanced diaminobenzidine (DAB) was used as the chromogen to produce a blue-black reaction product (35). GnRH was then demonstrated with a peroxidase-antiperoxidase procedure using DAB without enhancement to produce a brown reaction product. For the GnRH/dynorphin procedure, we used a polyclonal rabbit antibody to dynorphin A (1:10,000; 48 h at 4 C; Peninsula Laboratories, Inc., Belmont, CA, IHC 8730) and a mouse monoclonal antibody against GnRH (1:500; 48 h at 4 C; HU4H, gift of H. Urbanski, Oregon Primate Research Center, Beaverton, OR). For detection of ß-endorphin/GnRH, we used a mouse monoclonal antibody (1:500; 48 h at 4 C; Roche Molecular Biochemicals, Indianapolis, IN) and a rabbit polyclonal antibody (1:50,000; 48 h at 4 C; LR-1, gift from R. Benoit, Montreal General Hospital, Montreal, Canada ), respectively. Controls for double-labeling included omission of one or both primary antiserum, or preabsorption of diluted antiserum with nanomolar concentration of the appropriate corresponding peptide. Treatment of sections in this manner eliminated all specific staining.

Sections processed for light microscopic ICC were examined under bright-field illumination for occurrence of close associations between dynorphin or ß-endorphin varicosities and GnRH neurons in the diagonal band of Broca and medial septum (DBB), POA, anterior hypothalamus, ventrolateral hypothalamus, and MBH. Close appositions were defined as EOP-immunoreactive varicosities in direct contact with GnRH somas or dendrites viewed at the same plane of focus under high (x100) magnification. For each animal, the mean number of these close associations between EOP varicosities and each GnRH neuron was determined. In addition, the percentage of GnRH neurons in each region that possessed at least one close association, either dynorphin/GnRH or ß-endorphin/GnRH, was also calculated.

For EM visualization of EOP synapses onto GnRH neurons, we used a dual-label preembedding technique, in which the same close associations observed first at a light microscopic level can subsequently be examined at the ultrastructural level (36). The ICC procedure is similar to that described above, in that nickel-enhanced DAB and unenhanced DAB were used as labels for EOP and GnRH, respectively. The difference in the EM procedure is that 0.02% saponin (Sigma, cat. no. S1252) was used as a detergent for incubations in primary antibody rather than 0.1% Triton X-100, and detergent was eliminated from all other steps. For the GnRH/dynorphin procedure, we used a polyclonal rabbit antibody to dynorphin A (1:40,000; 48 h at 4 C; Peninsula Laboratories) and a mouse monoclonal antibody against GnRH (1:500; 48 h at 4 C; Sternberger Monoclonals Inc., Lutherville, MD). For detection of ß-endorphin/GnRH, we used a mouse monoclonal antibody (1:50; 48 h at 4 C; Roche Molecular Biochemicals) and a rabbit polyclonal antibody (1:10,000; 48 h at 4 C; LR-1, gift from R. Benoit), respectively. Controls for GnRH labeling using the mouse monoclonal antibody included omission of primary antiserum or preabsorption of diluted antiserum with nanomolar concentration of synthetic GnRH. Regions through the POA and MBH containing close associations between dynorphin or ß-endorphin immunoreactive fibers and GnRH cells were dissected out. Ten GnRH cells were analyzed in each region for dynorphin contacts and 10 GnRH cells from each region for ß-endorphin contacts. As a control for possible GnRH-GnRH contacts, we also stained alternate sections for GnRH alone and dissected out tissue containing GnRH perikarya in the POA and MBH. These tissue pieces were postfixed in 2% osmium tetroxide containing 1.5% potassium ferricyanide (37), dehydrated, and flat embedded in Epon 812. Semithin sections (1 µm) were cut and examined for the presence of GnRH cells and close contacts. Ultrathin (70 nm) sections were cut from the remaining block, mounted on formvar-coated grids, and stained with uranyl acetate and lead citrate. Sections were examined using a JEOL 1230 electron microscope (Japan Electron Optics, Ltd., Peabody, MA) and images of cells and terminals photographed at 4000, 10,000, and 20,000 magnifications.

Statistical analysis
LH pulses were identified using established criteria (5), and the effects of treatment in the POA or MBH on LH pulse frequency (pulses/4 h after implant insertion) were analyzed using Friedman’s two-way ANOVA. Mean LH concentrations were analyzed by one-way ANOVA with repeated measures and LH pulse amplitudes and latencies to the first LH pulse with one-way ANOVA (the absence of LH pulses in some ewes precluded a test using repeated measures). When significant differences were observed, Dunnett’s post hoc test was used to determine which groups differed from blank controls. The {chi}2 test was used to determine significant differences among groups in the proportion of ewes showing an increase in LH within three samples of implant insertion. Differences in EOP-close contacts on GnRH perikarya located in different areas were analyzed by one-way ANOVA. Statistical significance of P < 0.05 was used for all analyses.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Experiment 1. EOPs act via {kappa}-receptors in the MBH to inhibit LH pulse frequency
When ewes received empty microimplants into the MBH, LH pulse frequency was low, averaging slightly less than 1 pulse/4 h (Fig. 1Go) and the LH pulses that were observed occurred at variable times during the 4-h sampling period (Fig. 2Go and Table 1Go). As expected, implantation of naloxone produced an immediate increase in LH secretion (Fig. 2Go and Table 1Go) and significantly increased LH pulse frequency and mean LH concentrations over control values (Fig. 1Go). The patterns of LH secretion observed after implantation of the {delta}-antagonist were not significantly different from controls. In contrast, the {kappa}-antagonist produced an immediate LH pulse in five of the eight ewes (Table 1Go) and increased LH pulse frequency and mean LH levels to values similar to that seen with naloxone (Figs. 1Go and 2Go). The µ-antagonist appeared to produce increased LH secretion in some animals but not in others (Fig. 2Go), and the effect was often delayed so that the proportion of ewes showing an LH pulse shortly after microimplant insertion was not significantly different from that in ewes given empty implants (Table 1Go). Moreover, because this pattern was seen in only about half the ewes, no significant increase in LH pulse frequency or mean LH concentrations (Fig. 2Go) occurred with this treatment. There were no significant differences in LH pulse amplitudes among treatment groups (Table 1Go).



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  		FIG. 1. Antagonist to the {kappa}-receptor increases LH pulse frequency and mean LH concentrations when placed in the MBH of luteal phase ewes. Mean (+SEM) LH pulse frequencies (top panel) and LH concentrations (bottom panel) during the 4 h after insertion of blank (negative controls) tubing or tubing filled with naloxone (positive control) or specific antagonists to the {delta}-, {kappa}-, or µ-receptor into the MBH of luteal phase ewes (n = 8). *, P < 0.05 vs. negative control implants.

 


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  		FIG. 2. LH pulse patterns in response to microimplants of EOP antagonists into the MBH in two different ewes. Top panels, Empty (negative control) or naloxone-containing tubing (positive control) were inserted (arrows). Bottom panels, Tubing containing naltrindole ({delta}-antagonist), nor-BNI ({kappa}-antagonist), or naloxonazine (µ-antagonist) inserted as indicated by arrows. Solid circles, Peak of a statistically identified pulse.

 

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  		TABLE 1. Effect of EOP antagonists placed in to MBH

 
Implantation sites in all eight ewes were within approximately 1 mm of both the base of the brain and third ventricle and 1–3 mm anterior to the start of the pituitary stalk (Fig. 3Go, E–G). There was no obvious effect of anterior-posterior placements on the response to EOP antagonists. In two ewes, the microimplants were on the same side of the MBH. In these two ewes, the response to nor-BNI (1.0 ± 0 pulses/h) appeared to be less than in the ewes with bilateral implants (2.3 ± 0.4 pulses/4 h). However, the response to naloxone was similar in unilaterally and bilaterally implanted ewes (2.0 ± 1.0 vs. 1.7 ± 0.4 pulses/4 h).



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  		FIG. 3. Implantation sites in MBH and POA. Schematic representation of implantation sites in experiment 1 (bottom panels) and experiment 2 (top panels) shown on coronal sections of the ovine POA and hypothalamus. Implants in the same ewe are connected by a line. Solid circles indicate that nor-BNI and naloxone (hypothlamus) or nor-BNI, naloxonazine, and naloxone (POA) increased LH pulse frequency over controls. Open circles indicate placements at which these drugs had no effect; gray circles are placements at which only one of the drugs was stimulatory. Note the scale bar at lower right. ac, Anterior commissure; ARC, arcuate nucleus; f, fornix; LS, lateral septum; ME, median eminence; MPOA, medial preoptic area; MS, medial septum; mt, mammillothalamic tract; oc, optic chiasm; ot, optic tract; OVLT, organum vasculosum lamina terminalis; pt, pars tuberalis; SCN, suprachiasmatic nucleus; SON, supraoptic nucleus; st, stria terminalis; v, third ventricle; VMH, ventromedial hypothalamic nucleus.

 
Experiment 2. EOP act via both {kappa}- and µ-receptors in the POA to inhibit LH pulse frequency
Implantation sites in three of the 11 ewes were located rostral to the POA, in the anterior portion of the DBB (Fig. 3AGo); none of these ewes responded to naloxone, or other EOP antagonists, and data from them were excluded from analysis. Implants in the other eight ewes were centered in the medial POA in five ewes and just rostral or caudal to this area in the other three animals (Fig. 3Go, B–D). All microimplants were symmetrically bilateral and between 2 and 3 mm from midline and 1.5–2.5 mm from the base of the diencephalon.

LH pulse frequencies during control treatments were again low, averaging slightly more than one pulse per 4 h, and naloxone microimplants significantly increased LH pulse frequency and mean LH levels (Figs. 4Go and 5Go). Implantation of the {delta}-antagonist had no significant effect on LH secretion, but both the {kappa}- and µ-antagonists significantly increased LH pulse frequency and mean LH concentrations over control values (Fig. 5Go). The stimulatory action of these two receptor antagonists was observed in seven of the eight ewes and was similar in magnitude to that observed with naloxone. The response to the {kappa}-antagonist was more rapid than that to µ-antagonist (Fig. 4Go), based on the proportion of ewes with an increase in LH concentrations within 36 min of implantation (Table 2Go). There were no significant differences in LH pulse amplitudes among treatment groups (Table 2Go).



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  		FIG. 4. Antagonists to the {kappa}- and µ-receptor increase LH pulse frequency and mean LH levels when placed in the POA of luteal phase ewes. Mean (+SEM) LH pulse frequencies (top panel) and LH concentrations (bottom panel) during the 4 h after insertion of blank (negative controls) tubing or tubing filled with naloxone (positive control) or specific antagonists to the {delta}-, {kappa}-, or µ-receptor into the POA of luteal-phase ewes (n = 8). *, P < 0.05 vs. negative control implants.

 


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  		FIG. 5. LH pulse patterns in response to implants of EOP antagonists into the POA of two illustrative luteal-phase ewes. Top panels, Empty (negative control) or naloxone-containing tubing (positive control) were inserted (arrows). Bottom panels, Tubing containing naltrindole ({delta}-antagonist), nor-BNI ({kappa}-antagonist), or naloxonazine (µ-antagonist) inserted as indicated by arrows. Solid circles, Peak of a statistically identified pulse.

 

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  		TABLE 2. Effect of EOP antagonists placed in to POA

 
Experiment 3. Dynorphin- and ß-endorphin-containing neurons contact GnRH perikarya
Both dynorphin- and ß-endorphin-containing varicosities appeared to be closely associated with GnRH perikarya in all regions examined (Fig. 6Go). An average of 488 and 516 GnRH neurons/ewe were examined for dynorphin and ß-endorphin close associations, respectively, and their anatomical distribution was the same as previously observed (25, 26). However, there was a regional difference in the dynorphin input. Almost 90% of the GnRH perikarya in the MBH received at least one dynorphin close association, compared with only about 40% of the GnRH perikarya in other areas, and the mean number of dynorphin boutons close to GnRH perikarya in the MBH was twice the number on GnRH perikarya in other areas (Fig. 7Go). In contrast, no regional differences were observed in the percentage of GnRH neurons with ß-endorphin-positive close associations. The mean number of ß-endorphin boutons close to GnRH perikarya in the MBH was not greater than those on GnRH perikarya in the POA but was significantly different from mean number of inputs on GnRH perikarya in other areas (Fig. 7Go).



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  		FIG. 6. Close associations between ß-endorphin (A) or dynorphin (B) varicosities and GnRH neurons in the POA (A) and MBH (B) as seen in dual immunoperoxidase-labeled sections. Black arrows, Close contacts between ß-endorphin (A) or dynorphin (B) varicosities (blue-black) and GnRH cells (brown). In B, dynorphin-positive cell bodies (blue-black) are also seen. Bar, 25 µm.

 


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  		FIG. 7. Dynorphin-containing (top panel) and ß-endorphin-containing (bottom panel) boutons contacting GnRH perikarya in the MBH and POA of luteal-phase ewes (n = 3). Mean (+SEM) percentage of GnRH neurons contacted by at least one EOP-positive varicosity (solid bars) and number of EOP-positive close contacts per neuron (stippled bars) in the DBB, POA, anterior hypothalamic area (AHA), ventrolateral hypothalamus (VLH), and MBH. *, Significantly different (P < 0.05) from all other bars (top panel); bars with different superscripts are significantly different in bottom panel.

 
Experiment 4. Both dynorphin and ß-endorphin neurons synapse on GnRH cells
Because close associations observed at the light microscopic level do not necessarily reflect synaptic input, we next determined whether terminals containing these two EOPs synapse directly onto GnRH perikarya. Dynorphin- and ß-endorphin-positive axon terminals were abundant in the ovine POA and MBH and contained immunoreactive dense core vesicles. Dynorphin-containing boutons, seen at a light microscopic level to be in close contact with GnRH perikarya, formed asymmetric synapses onto GnRH somas and dendrites when observed at an ultrastructural level (Fig. 8Go). Similarly, light microscopic close contacts between ß-endorphin varicosities and GnRH neurons were also confirmed to be synaptic inputs at an EM level (Fig. 9Go). Every EOPcontaining close association identified at the light level that was also examined at the EM level was observed to be either a synapse or direct membrane contact. In control sections through the POA and MBH that were single labeled for GnRH, we saw no instances of immunoreactive terminals contacting immunoreactive somas or dendrites.



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  		FIG. 8. Dynorphin synaptic input onto a GnRH dendrite. A, Low-power electron micrograph of a dynorphin-positive terminal (arrow), which synapses onto the dendrite (d) of a MBH GnRH neuron. Bar, 1 µm. B, High-power electron micrograph of the same terminal showing the presence of a well-defined synaptic cleft (arrows) and asymmetric synaptic density. The terminal contains an immunoreactive dense core vesicle (d) as well as unlabeled mitochondria (m). ax, Myelinated axon.

 


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  		FIG. 9. ß-Endorphin synaptic input onto a GnRH soma. A, Low-power electron micrograph of a ß-endorphin-containing axon terminal (white asterisk) contacting the soma of a POA GnRH neuron. Nonimmunoreactive axon terminals (e.g. black asterisk) can also be seen contacting this soma. nc, Nucleus. Bar, 2 µm. B, High-power electron micrograph of the same terminal showing the presence of a synaptic density (arrow) and immunoreactive dense core vesicles (e.g. arrowheads) within the terminal.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The results of this study provide strong evidence that dynorphin and ß-endorphin inhibit GnRH pulse frequency in luteal phase ewes and raise the possibility that this action occurs via direct synaptic input to GnRH perikarya. The electron microscopic analyses clearly demonstrate that both dynorphin- and ß-endorphin-containing axon terminals synapse upon GnRH neurons. The latter has been observed in the rhesus monkey (38) and rat (39), but this is the first report of direct dynorphin input to GnRH cells. Although these observations are consistent with direct EOP inhibition of GnRH release, it should be noted that although rodent GnRH neurons receive ß-endorphin input (39), they apparently do not contain any of the known EOP receptors, based on the lack of in situ hybridization for the appropriate mRNAs (40, 41). Furthermore, pharmacological evidence in the rat suggests that EOPs influence GnRH secretion indirectly via presynaptic inhibition of norepinephrine (42). We are currently determining whether ovine GnRH neurons contain EOP receptors and are reluctant to conclude that the EOP synapses observed here are functionally important until this issue is resolved.

Assessment of the effects of the microimplants is based on two important considerations: the spread of the drugs from the site of implantation and the specificity of the antagonists used. We could not determine the volume of tissue affected by the microimplants, but the spread of radioactivity from similar crystalline implants is limited to a radius of about 1 mm (43, 44, 45). The ineffectiveness of three different antagonists, implanted just rostral to the POA in which they were effective (Fig. 3Go), suggests that these drugs are also limited in their diffusion. Based primarily on in vitro assessments, the three specific EOP receptor antagonists used have affinities for their specific receptor 100–1000 times their affinity for the other two EOP receptors (29, 30). Thus, we have based our interpretations on the assumption that each drug is acting locally at its targeted receptor.

Because it is generally recognized that dynorphin, but not other EOPs, acts via {kappa}-receptors (46, 47), the consistent ability of the specific {kappa}-antagonist, nor-BNI, to increase LH pulse frequency indicates that this EOP holds GnRH pulse frequency in check during the luteal phase of the ovine estrous cycle. Moreover, the inability of selective µ- and {delta}-receptor antagonists to significantly increase LH when placed into the MBH supports the hypothesis that dynorphin is the only EOP acting in this area. In contrast to the data from MBH implants, both µ- and {kappa}-receptor antagonists increased episodic LH release when placed in the POA, suggesting that another EOP is also acting in this area. Based on immunoneutralization studies, this second EOP is most likely ßendorphin (23). Data in the rat are also consistent with inhibition of GnRH pulses by both dynorphin acting via {kappa}-receptors (21, 48) and ß-endorphin acting via µ-receptors (19, 20), but most workers appear to agree that in the rat opioid effects are generally believed to involve ß-endorphin acting on the µ-receptor (14). The present data suggest that either there are species differences in the EOP control of LH pulse frequency or that the weight of evidence in the rat needs to be reevaluated.

In interpreting the effects of these EOP antagonists, we assumed that they are disrupting the negative feedback action of progesterone for two reasons. First, they selectively increased LH pulse frequency, the characteristic of episodic LH secretion that is controlled by progesterone (2, 3, 4, 5). If the antagonists were affecting estradiol-negative feedback, they should have increased LH pulse amplitude (2, 8, 9). Second, there is extensive evidence from systemic administration of EOP antagonists that EOPs mediate progesterone-negative feedback in the ewe (49). For example, EOP antagonists, such as naloxone, increase LH pulse frequency in progesterone-treated OVX ewes but do not alter pulse frequency in estradiol-treated OVX (9, 15, 16, 17) or OVX ewes not treated with steroids (9, 16).

Whereas effects of the microimplants confirm previous data that EOPs act in both the MBH and POA to hold GnRH pulses in check during the luteal phase (23, 27), the high density of dynorphin-positive boutons seen specifically close to GnRH perikarya in the MBH (Fig. 7Go) raises the possibility that dynorphin inhibition of this subset of GnRH cells is particularly important for progesterone-negative feedback of LH pulse frequency. This concept is supported by two other lines of evidence. First, systemic administration of a nonspecific EOP antagonist to luteal phase ewes induced Fos expression in GnRH neurons within the MBH but not in GnRH neurons outside this area (26). Second, knife cuts between the POA and MBH that sever most connections between these areas do not alter the luteal phase of the ovine estrous cycle (50) and do not affect the EOP system mediating progesterone-negative feedback (51). The latter observation also points to EOP perikarya within the MBH as critical to this action of progesterone. Consistent with this hypothesis is recent evidence that a very high percentage of dynorphin cells (>90%) in the arcuate nucleus of the ewe contain progesterone receptors (52). Based on these observations, our current working hypothesis is that dynorphin-GnRH interactions within the MBH are critical elements for progesterone inhibition of LH pulse frequency and the actions of EOPs in the POA provide redundancy to this system and/or mediate other actions of progesterone or estradiol important for the control of the GnRH surge.

Whereas this study has focused primarily on control of pulsatile GnRH secretion by progesterone-negative feedback, it is important to keep in mind that the anatomical data on EOP inputs to the GnRH perikarya may also be relevant to EOP influences on the preovulatory GnRH surge. Because the ßendorphin/µ-receptor system has been implicated in control of the preovulatory surge (12, 13, 24), the observations of similar ß-endorphin input on to GnRH neurons regardless of location is particularly relevant. These data are consistent with a recent confocal microscopic analysis in sheep (53), which also observed three to four ß-endorphin-positive close contacts on GnRH soma regardless of anatomical location. The data raise the possibility that removal of ß-endorphin inhibition on GnRH perikarya throughout their rostral-caudal continuum is permissive for the preovulatory GnRH surge. This hypothesis is also consistent with reports that: 1) GnRH cells that express Fos during the surge are equally distributed among all POAhypothalamic areas (54); and 2) a µ-agonist can delay the LH surge when administered locally to the ovine POA or MBH (55). In light of these anatomical, pharmacological, and physiological data, our working hypothesis for EOP control of GnRH secretion in the ewe is that the dynorphin/{kappa}-receptor system mediates progesterone-negative feedback, whereas the ß-endorphin/µ-receptor system primarily influences the timing of preovulatory GnRH surge. It should be noted, however, that opioid antagonists do not advance the timing of the LH surge in the ewe (56, 57) so that the specific role, if any, of the EOP system in the stimulatory effects of estradiol (56, 57) or the inhibitory effects of progesterone (58, 59) on the GnRH surge in sheep remains to be determined.

Although any inferences on the role of the dynorphin/{kappa}-receptor in progesterone-negative feedback from the observations reported here are obviously limited to the ewe, they also may be relevant to other species. In the rat, there is strong evidence that {kappa}-receptors mediate the inhibition of pulsatile LH secretion during pregnancy (21) and that this inhibition occurs in both the POA and MBH (48). The role of {kappa}-receptors in mediating progesterone-negative feedback has not been examined in other species, but it is interesting to note that EOPs have been implicated in the inhibition of LH secretion during the luteal phase in a number of other species including pigs (60), rhesus monkeys (4, 7), and humans (7, 61).

In summary, the results of this study, together with the recent demonstration that a very high percentage of dynorphin neurons contain nuclear progesterone receptors (52), strongly suggest that the dynorphin/{kappa}-receptor system plays a critical role in mediating progesterone-negative feedback in the ewe. Observations of dynorphin inputs also raise the possibility that this inhibition may be exerted directly on GnRH perikarya within the MBH. Dynorphin and ß-endorphin inputs to GnRH neurons in the POA may provide redundancy to progesterone-negative feedback, may be involved in other actions of progesterone or estradiol important for the control of the GnRH surge, or may mediate the effects of other internal or external signals on GnRH secretion.


   Acknowledgments
 
We thank Drs. Gordon Niswender and Dr. Al Parlow and the National Pituitary Agency for reagents used in RIA, Paul Harton and Bob McTaggert for histological assistance, and Sarah Beemer and Kerie Miller for animal care.


   Footnotes
 
This work was supported by grants from NIH (HD17864, HD39916) and the U.S. Department of Agriculture (2000-02132).

Preliminary reports of these experiments were presented at the 32nd Annual Meeting of the Society for the Study of Reproduction, Pullman, Washington, 1999, and 29th Annual Meeting of the Society for Neuroscience, Miami, Florida, 1999.

Current address for G.M.A.: Department of Anatomy and Structural Biology and Centre for Neuroendocrinology, University of Otago Medical School, Dunedin 9001, New Zealand.

Current address for S.L.H.: University of Pittsburgh Medical Center, Department of Anesthesiology, University of Pittsburgh, 1311 Biomedical Sciences Tower, Pittsburgh, Pennsylvania 15261.

Abbreviations: DAB, Diaminobenzidine; DBB, diagonal band of Broca; EM, electron microscopic; EOP, endogenous opioid peptide; ICC, immunocytochemistry; MBH, medial basal hypothalamus; nor-BNI, nor-binaltorphimine; OVX, ovariectomized; PB, phosphate buffer; POA, preoptic area.

Received September 29, 2003.

Accepted for publication February 17, 2004.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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