Progesterone Increases Dynorphin A Concentrations in Cerebrospinal Fluid and Preprodynorphin Messenger Ribonucleic Acid Levels in a Subset o
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内分泌学杂志 2005年第4期
Department of Cell Biology, Neurobiology, and Anatomy (C.D.F., M.N.L.), University of Cincinnati College of Medicine, and Neuroscience Program (C.D.F., M.N.L.), University of Cincinnati, Cincinnati, Ohio 45267-0521; and Department of Physiology (R.L.G., V.L.A., M.V.), West Virginia University Health Sciences Center, Morgantown, West Virginia 26506-9229
Address all correspondence and requests for reprints to: Dr. Michael N. Lehman, Department of Cell Biology, Neurobiology, and Anatomy, University of Cincinnati College of Medicine, Cincinnati, Ohio 45267-0521. E-mail: michael.lehman@uc.edu.
Abstract
Recent studies suggest that the endogenous opioid peptide, dynorphin, is an important mediator of progesterone negative feedback on GnRH pulse frequency in the ewe. These experiments tested this hypothesis by examining the effects of progesterone on dynorphin A concentrations in cerebrospinal fluid (CSF) collected from the third ventricle and expression of preprodynorphin (PPD) mRNA in hypothalamic nuclei. CSF was collected every 10 min for 5 h in three groups of ewes: 1) ovary-intact ewes during the luteal phase (d 6–7 of estrous cycle); 2) ewes 6–7 d after ovariectomy (OVX); and 3) OVX ewes treated for 6–7 d with implants that produced luteal-phase progesterone levels. Diencephalic tissue from these ewes was then collected and processed for in situ hybridization using an ovine cDNA probe against PPD. Progesterone treatment increased dynorphin A concentrations in CSF over that observed in untreated OVX ewes; CSF dynorphin A concentrations in ovary-intact ewes were midway between the other groups. OVX significantly decreased the number of PPD mRNA-expressing cells in the preoptic area (POA), anterior hypothalamic area (AHA), and arcuate nucleus (ARC), with no change seen in any other PPD-expressing nuclei. Progesterone treatment of OVX ewes restored PPD expression in the POA and AHA to levels seen in luteal-phase animals but had no effect on PPD expression in the ARC. These results are consistent with the hypothesis that progesterone acts via dynorphin neurons to inhibit pulsatile GnRH secretion and point to dynorphin neurons in the POA, AHA, and ARC as potential mediators of this action during the luteal phase.
Introduction
THE SECRETION OF GnRH into the pituitary portal blood supply is the final common pathway responsible for neuroendocrine control of reproduction. The activity of GnRH neurons, in turn, is controlled by the positive and negative feedback actions of estradiol and progesterone (1, 2, 3, 4). Specifically, elevated estradiol triggers the preovulatory GnRH surge at the end of the follicular phase, while estradiol and progesterone act singly, and in concert, to inhibit the episodic secretion of GnRH that occurs throughout the rest of the cycle.
Although both estradiol and progesterone inhibit GnRH pulses, they do so by different mechanisms. Estradiol alone inhibits LH (see Ref. 31) and GnRH (see Ref. 43) pulse amplitude, whereas progesterone by itself inhibits LH (see Ref. 31) and GnRH (5) pulse frequency, an action that is enhanced by estradiol (see Ref. 40). In sheep, this action of progesterone appears to be mediated by the classical nuclear progesterone receptor (PR) (5), but PR are generally not found in GnRH neurons in this and other species (6, 7, 8, 9). Thus, other systems most likely transmit the actions of progesterone to GnRH neurons. There is now general agreement that endogenous opioid peptides (EOP) are one of the major systems that mediate progesterone negative feedback in sheep (1, 10), rats (11), and primates (2, 12, 13). In contrast, there is clear evidence that EOP do not mediate the inhibition of GnRH pulse amplitude by estradiol in sheep (see Ref. 44), and there is some evidence implicating noradrenergic neurons in this action of estradiol (see Ref. 45).
Recent work has raised the possibility that dynorphin A1–17 may be the primary EOP involved in progesterone negative feedback. Dynorphin A1–17 is part of a family of EOP derived from preprodynorphin (PPD) that also includes dynorphin A1–8 and dynorphin B. Specifically, three lines of evidence have implicated dynorphin A, which is the endogenous ligand for the EOP receptor (see Ref. 46): 1) almost all GnRH neurons in the medial basal hypothalamus, and about 50% of those elsewhere in the sheep brain, receive synaptic inputs from dynorphin A-containing terminals (14); 2) an EOP receptor antagonist consistently increases LH pulse frequency in luteal phase ewes (14); and 3) over 90% of the dynorphin A-positive perikarya in the preoptic area (POA), anterior hypothalamus (AHA), and arcuate nucleus (ARC) contain PR (15).
If this working hypothesis is correct, then progesterone should increase dynorphin release from the synapses that terminate on GnRH neurons. However, this prediction is difficult to test directly because of the dispersed nature of GnRH perikarya (16). Therefore, we have chosen two less-direct approaches to testing it. First, we assessed the effect of progesterone on dynorphin A concentrations in the cerebrospinal fluid (CSF) collected from the third ventricle (IIIV); third ventricular CSF GnRH levels in the sheep correlate well with its endogenous secretion rates (17). Second, we tested whether progesterone would increase PPD mRNA levels in distinct subpopulations of dynorphin-containing neurons, an approach that has been useful in assessing the effects of steroids on other EOP in the sheep (18, 19) and rat (20, 21, 22, 23).
Although there are similarities in the EOP mediation of progesterone negative feedback in sheep, rats, and primates, there is one difference in this action of progesterone among these species. In primates (2) and rats (24), the negative feedback action of progesterone requires the presence of estradiol; whereas in sheep, progesterone alone inhibits tonic LH secretion in acutely ovariectomized (OVX) animals (25). Consequently, many studies in sheep have examined this action of progesterone in the absence of estradiol (1, 2, 5, 10, 31); and there is evidence that, by itself, progesterone can affect the expression of the mRNA for EOP in sheep (18, 19). Because progesterone negative feedback does not require estradiol and estradiol negative feedback does not involve EOP, in this study we examined the actions of progesterone on dynorphin A concentrations in CSF and PPD mRNA levels in the absence of estradiol.
Materials and Methods
Animals
Adult Suffolk ewes were maintained in an open barn with free access to water and fed once daily with a maintenance regimen of silage supplemented with grain. This study was performed during the breeding season (January and early February) in 23 ewes that had demonstrated at least two normal 16- to 17-d estrous cycles (determined by monitoring estrous behavior with a vasectomized ram). They were moved to indoor facilities 2–3 d before experimentation. Once indoors, the animals were housed two ewes/pen under a photoperiod similar to that occurring outdoors. The routine handling and experimental procedures involving animals were approved by the West Virginia University Animal Care and Use Committee.
Experimental protocols
Ovarian cycles were synchronized by injection of prostaglandin F2, 7 d apart (26). Three days after the second prostaglandin F2 injection (approximately d 0 of next ovarian cycle), ewes were either OVX (n = 16) or sham OVX (n = 7) and a cannula inserted into the IIIV. Eight of the OVX ewes received two SILASTIC-brand (Dow Corning Corp., Midland, MI) packets of progesterone placed sc just after OVX to produce midluteal phase P concentrations (27). Blood and CSF samples were then collected 6–7 d after surgery. There were thus three treatment groups: ovary-intact (n = 7) on d 6 or 7 of the estrous cycle, OVX (n = 8), and OVX+P (n = 8).
Surgeries
Ovariectomies were preformed by midventral laparotomy using sterile procedures under gas anesthesia (oxygen+nitrous oxide, supplemented with halothane as needed). After OVX (or sham OVX, which consisted of midventral incision), a 16-gauge stainless steel cannula was placed into the IIIV using a stereotaxic procedure as previously described in detail (28). Briefly, the surface of the skull was exposed, a 1-cm-diameter hole drilled, and the sagittal sinus ligated and cut. Radio-opaque dye was then injected into one lateral ventricle, and frontal and lateral x-rays were used to guide the placement of the cannula with its tip about 5 mm dorsal to the median eminence. Placement in the IIIV was confirmed by flow, or withdrawal of CSF, from the cannula. The cannula was then cemented to the skull with screws and dental acrylic, plugged, and covered with a protective cap. All ewes were monitored closely post operation, and only those that showed complete recovery (based on temperature, food intake, and level of arousal) within 3 d were used in these experiments.
Blood and CSF collections
Collection of CSF was based on the procedure described by Skinner et al. (29). The day before collection, one jugular vein was catheterized and a panel used to restrict each animal’s movement so that it could lie down but not turn around. The next morning, polyethylene tubing (0.38-mm inside diameter, 1.09-mm outside diameter; VWR International, Bridgeport NJ) was inserted down the cannula so that it extended about 1 mm beyond its end into the ventricle. This tubing was connected via silicone rubber tubing (1.0-mm inside diameter, 2.0-mm outside diameter; Sil-Med Corp., Taunton, MA) to a Gilson peristaltic pump, and samples were collected every 10 min for 5 h. Approximately 0.5 ml CSF was collected on ice continuously over the 10-min period and was immediately frozen after collection. Heparinized blood samples (5 ml) were taken every 10 min, from the catheter previously placed into the jugular vein, for 5 h. In a few cases, flow through the peristalic pump failed and samples were continued to be collected directly from the cannula with a syringe every 10 min. In six ewes, only a few CSF samples could be collected; these data were not included in the analyses, leaving five to six animals/group. Plasma was stored at –20 C and CSF samples stored at –80 C until assayed.
Tissue collection
The day after CSF and blood sample collection, fresh tissue was collected for in situ hybridization analysis of PPD mRNA expression. Animals were given an anesthetic dose of sodium pentobarbital (1 g, iv), and the protective cap and cannula were removed to facilitate rapid collection of tissue. The ewes were then given a lethal dose of pentobarbital (2 g, iv) and decapitated; and a tissue block containing the septal region, POA, and hypothalamus was rapidly dissected out and placed on dry ice. After freezing, the blocks were shipped on dry ice to the University of Cincinnati and stored at –80 C until sectioned. Coronal sections (20 μm) were cut on a cryostat, thaw mounted on SuperFrost plus slides (Fisher Scientific, Pittsburgh, PA), and stored at –80 C until processed for in situ hybridization.
RIAs
LH was measured in duplicate aliquots of all plasma samples using a previously described RIA (27), except that we iodinated-purified ovine LH provided by Dr. Parlow and the National Hormone and Peptide Program for tracer. Assay sensitivity averaged 0.18 ng/ml (NIH-S24). Intraassay and interassay coefficients of variation (CVs) averaged 9% and 13%, respectively. Progesterone was measured in 100-μl aliquots of plasma samples collected at three time points spanning the 5-h collection period using a commercially available kit (Diagnostic Systems Laboratories, Inc., Webster, TX) previously validated for use in sheep (28). All samples were measured in a single assay, with a sensitivity of 0.02 ng/ml and intraassay CV of 9%.
Dynorphin concentrations in CSF were measured using a commercial kit (Peninsula Laboratories, Inc., Belmont, CA). Briefly, 100 μl rabbit dynorphin A antisera was added to the standard curve (in triplicate, ranging from 1–32 pg) and duplicate aliquots of CSF (usually 150 μl) that had been lyophilized and reconstituted in 100 μl assay buffer. After an overnight incubation at 4 C, iodinated-dynorphin A (100 μl; 15,000 cpm) was added, and incubation at 4 C continued for another 24 h. Antibody-bound radioactivity was separated from unbound with donkey antirabbit -globulin on d 3. This assay has minimal cross-reactivity (<1%) with dynorphin A (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13) and no cross-reactivity with dynorphin B, ?-endorphin, or leu-enkephalin. The assay sensitivity averaged 1.0 ± 0.2 pg (n = 6), and the ED50 was 12.1 ± 0.6 pg (n = 6). Pools of CSF from three different ewes showed parallelism with the standard curve (Fig. 1). Almost all (>95%) unknown samples fell between 80% and 50% displacement of iodinated dynorphin, and the intraassay CVs of variation at these points on the standard curve were 19 ± 0.8% and 12.1 ± 1.1%, respectively; interassay CV averaged 12% for the six assays in which CSF levels were monitored.
FIG. 1. Displacement of iodinated-dynorphin from dynorphin-A antibody by 1–32 pg dynorphin (lower plot and x-axis) and 50, 100, and 200 μl of three pools of CSF (top axis and upper plots).
In situ hybridization
Probes were produced using a cDNA sequence of a portion of the ovine PPD previously described (30). Antisense and sense RNA fragments were generated from linearized templates (2 μg) using T7 and SP6 RNA polymerases (Promega, Madison, WI), respectively, in a transcription reaction containing: 5x transcription buffer (Promega), 5 mM ribonucleotide triphosphates (GTP, CTP, ATP), 100 μM UTP, ribonuclease (RNase) inhibitor (Rnasin, Promega, N211A), 100 mM dithiothreitol, and 100 μCi 35S-UTP (3000 Ci/mmol; NEN Life Science Products, Boston, MA) for 2 h at 37 C. Samples were then incubated with Dnase-I (Sigma, St. Louis, MO) at 37 C for 15 min to remove template. The reaction was stopped with 0.5 EDTA. Unincorporated isotopes were removed using spin columns (Roche, Indianapolis, IN). Probes were diluted in Tris, EDTA, containing 10 mg/ml Torula RNA (Sigma), denatured at 90 C for 3 min, and combined with hybridization buffer containing 50x Reinhardt’s solution, deionized formamide, 50% dextran sulfate, 5 M NaCl, 1 M Tris (pH 8.0), 0.5 EDTA, and 1 M dithiothreitol.
Just before use, slides containing tissue sections were removed from the freezer, placed in racks, and dried for 30–40 sec. Sections were then fixed for 5 min in 4% paraformaldehyde in phosphate buffer (PB, pH 7.4) at room temperature and then rinsed twice in PB, while stirring. Slides were dipped briefly in water, then in triethanolamine, and finally, soaked in triethanolamine plus acetic anhydride for 10 min, while stirring. Slides were then soaked in 2x saline sodium citrate (SSC: 0.15 M NaCl, 0.015 M sodium citrate); 70%, 95%, and 100% ethanol, respectively; followed by delipidation in chloroform for 5 min. Slides were soaked for 3 min each in 100% and 95% ethanol and then air-dried. Once dried, hybridization buffer with labeled probe (1 x 106 cpm/slide) was applied to the slides in a vol of 150 μl. Calculations based on molar amount of probe and estimates of tissue RNA indicated that this probe is more than 1000-fold in excess of endogenous levels. Slides were coverslipped with Parafilm (Fisher Scientific) and sealed with rubber cement and hybridized overnight at 46 C. On the following day, coverslips were removed, and the slides were washed for 15 min in 4x SSC. Slides were then incubated for 30 min at 37 C in RNase buffer (1 M Tris, pH 8.0; 5 M NaCl; 0.5 M EDTA) containing RNase-A (0.03 mg/ml; Sigma) followed by incubation for 30 min at 37 C in buffer without RNase-A. Next, the slides were washed at room temperature under increasingly stringent conditions (2x SSC, 0.1x SSC [59 C, twice, 30 min each], 0.1 x SSC [room temperature, twice]). Finally, slides were dehydrated for 3 min each in 50, 85, and 100% ethanol and then air-dried. Slides were then dipped in Kodak NTB3 emulsion, dried, and exposed in the dark for 21 d at 4 C. The emulsion was developed in Dektol (Kodak, Rochester, NY). The sections were counterstained with cresyl violet, dehydrated, and coverslipped with DPX (EM Sciences, Fort Washington, PA). Sense controls and RNase-pretreated sections showed only nonspecific labeling and background levels comparable with antisense-treated sections.
Data analysis
Number of cells expressing PPD.
Sections were coded so that analysis would be preformed blind to the condition of the animals. The locations of PPD-expressing cells were identified in the POA and hypothalamus of each ewe under dark-field and bright-field microscopy using a Leica microscope. Every tenth section was analyzed. Regions chosen for analysis were those shown to contain large numbers of PPD mRNAexpressing cells in our previous study (30): the POA, AHA, bed nucleus of the stria terminalis (BNST), supraoptic nucleus (SON), paraventricular nucleus (PVN), ventromedial nucleus (VMN), dorsomedial nucleus of the hypothalamus (DMH), and ARC. Three sections were analyzed for each area. Individual circular deposits of silver grains were considered positive for PPD if grains were 5x over background density. Images of labeled material were captured using a digital camera (Magnafire; Optronics, Goleta, CA) attached to a Leica microscope (Deerfield, IL) and imported into Adobe Photoshop 7.0 (Adobe Systems, San Jose, CA). Images were not altered in any way except for minor adjustments of brightness and contrast. Low-power drawings of the sections analyzed were made with a camera lucida to help provide orientation for the dark-field images.
Percent of nucleus covered by grains.
For every area analyzed, 12 cells with clear, Nissl-stained nuclei were randomly chosen, and high-powered (x40) bright-field photomicrographs were taken of each (Fig. 2). Because Nissl staining of the cytosol was uneven, it was impossible to determine whether grains overlying regions outside the stained nuclei were due to the cell in question or neighboring cells. Thus, we chose a more conservative quantitation method and analyzed silver grains over the nucleus. Using NIH Image, the area of the Nissl-stained nucleus of each cell was outlined (e.g. Fig. 2), and the total area of the nucleus was found. A standard threshold was chosen so that no pixels were attributed to the Nissl-stained nucleus and only pixels associated with silver grains were measured. The total number of silver grain pixels within the area of the nucleus was determined. Then the same shape traced around the nucleus was used to analyze an area lacking Nissl staining, and the total number of pixels was measured within this area. This created a background pixel number. The total number of pixels attributed to background silver grain deposits was subtracted from the number of pixels of the silver grains over the nucleus. This number was then divided by the total area outlined by the nucleus to arrive at the percent of the nucleus covered by grains. The percent grain coverage for the 12 random cells was averaged to obtain a mean percentage of the nucleus covered by grains for each area in each animal.
FIG. 2. High-power bright-field photomicrograph showing an example of a PPD-positive cell in the ARC. Silver grains are seen overlying the blue, Nissl-stained nucleus of this cell (outlined area) as well as surrounding it. Scale bar, 10 μm.
Statistical analysis
Hormones and CSF dynorphin concentrations.
LH pulses were identified using previously published criteria (31) and differences among treatment groups in LH pulse frequency evaluated by ANOVA. Mean differences in dynorphin concentrations in CSF and P in serum, the number of cells expressing PPD, and the average percent of nucleus covered by grains among treatment groups were all analyzed by one-way ANOVA followed by Tukey’s post hoc test. Changes in serum progesterone and CSF dynorphin concentrations over the 5-h period of collection, within each treatment group, were analyzed by one-way ANOVA with repeated measure and linear regression analysis, respectively. P < 0.05 was considered statistically significant for all analyses.
Results
Serum hormone and CSF dynorphin concentrations
As expected, there was no difference in the peripheral progesterone concentrations between the intact and OVX+P groups, whereas the OVX ewes showed levels of progesterone at the lower limit of detection (Fig. 3). LH pulse frequency in the ovary-intact and OVX+P groups averaged approximately 1 pulse/5 h and was significantly increased to 4 pulses/5 h in the OVX group (Figs. 3 and 4).
FIG. 3. Histogram comparing the concentration of CSF dynorphin, serum levels of progesterone, and LH pulse frequency (FREQ) in intact luteal controls (Intact, n = 6), OVX (n = 5), and OVX ewes bearing progesterone implants (OVX+P, n = 6). Different letters signify P < 0.05.
FIG. 4. Concentrations of plasma LH (bottom panels) and of dynorphin in CSF collected from the IIIV (top panels) of an ovary-intact ewe (left panels), an OVX (middle panels) ewe, and an OVX ewe treated with luteal-phase P levels (right panels). Solid circles in plots of LH indicate peaks of LH pulses.
Dynorphin A concentrations in CSF collected from the IIIV showed marked temporal variations, some as great as 50–100% over a 10- to 20-min sampling period, suggestive of an episodic mode of release (Fig. 4). However, there was no obvious correlation between the patterns of dynorphin in CSF and LH pulses in peripheral circulation or progesterone treatment (Fig. 4). The OVX ewes treated with progesterone had a significantly higher mean dynorphin concentration in CSF compared with that in OVX animals (Fig. 3). Intact ewes had intermediate CSF dynorphin levels and did not differ significantly from levels in either the OVX or OVX+P groups.
In OVX and OVX+P animals, peripheral progesterone concentrations were relatively constant during the 5-h collection period; but in the ovary-intact ewes, progesterone levels significantly increased from 1.4 ± 0.2 to 2.2 ± 0.3 ng/ml over this period (Fig. 5). During this same period, there was a gradual increase in CSF dynorphin concentrations in ovary-intact ewes (Fig. 5) that was statistically significant (P < 0.001) by linear regression analysis. There were no significant trends over the 5-h collection period in CSF dynorphin concentrations in the OVX or OVX+P groups (data not shown).
FIG. 5. Mean (±SEM) serum progesterone (PROG) concentrations (bars) and dynorphin concentrations (solid circles) in CSF samples collected from ovary-intact ewes during the luteal phase of the estrous cycle (n = 6).
Expression of PPD mRNA
As previously described (30), PPD mRNA-expressing cells were found in a number of nuclei of the ovine POA and medial basal hypothalamus, including the POA, AHA, BNST, SON, PVN, VMN, DMH, and ARC. Three of these areas, the POA (Fig. 6, A and B), AHA (Fig. 6, D–F), and ARC (Fig. 7), showed a significant change in the number of cells expressing PPD mRNA among the three experimental groups (see Fig. 10). Other areas, including the SON (Fig. 8), PVN (Fig. 9), VMN, DMH, and BNST, showed no significant differences among groups in the number of cells expressing PPD mRNA (Fig. 10).
FIG. 6. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the ventromedial POA (A–C) and AHA (D–E) of intact luteal ewes (A and D), OVX ewes (B and E), and OVX ewes bearing implants that maintain luteal phase levels of progesterone (OVX+P) (C and F). For orientation, camera lucida drawings of the sections from the intact ewes (A and D) are displayed at left, and the solid boxes within these drawings show the areas depicted in the photomicrographs. Note that the number of labeled cells in the rostral portion of the PVN (rPVN) (arrows in D and E) does not change. OVLT, Organum vasculosum of the lamina terminalis; 3v, IIIV; ac, anterior commissure; oc, optic chiasm. Scale bar, 200 μm.
FIG. 7. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the ARC of intact luteal phase (A), OVX (B), and OVX+P (C) ewes. The boxed area in the camera lucida drawing at left shows the region depicted in the photomicrographs. fx, Fornix; mt, mammillothalamic tract. Scale bar, 200 μm.
FIG. 10. Histogram comparing the mean (±SEM) number of PPD mRNA-expressing cells in the POA, AHA, ARC, PVN, VMN, DMH of luteal phase (Intact, n = 7), OVX (n = 8), and OVX ewes bearing progesterone implants (+P, n = 8). Asterisks, P < 0.05.
FIG. 8. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the SON (A–C) of intact luteal controls (A), OVX ewes (B), and OVX+P ewes (C). The boxed area in the camera lucida drawing at left shows the region depicted in the photomicrographs. Scale bar, 200 μm.
FIG. 9. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the PVN (A–C) of intact luteal controls (A), OVX ewes (B), and OVX+P ewes (C). The boxed area in the camera lucida drawing at left shows the region depicted in the photomicrographs. ot, Optic tract. Scale bar, 200 μm.
Compared with intact animals, OVX ewes showed almost a 50% decrease in the number of PPD mRNA-expressing cells in the POA and AHA and a 30% decrease in the ARC (Figs. 6, A–C, and 10). Progesterone treatment of OVX ewes increased the number of PPD mRNA-expressing cells in the POA and AHA to levels not significantly different from intact controls (Figs. 6 and 10). However, in the ARC, the number of cells expressing PPD mRNA in the P-treated group did not return to the intact level, and remained at the OVX level of expression (Figs. 7, A–C, and 10). Neither OVX nor progesterone treatment of OVX ewes affected the percentage of the cell nucleus covered by silver grains for any of the areas analyzed (Table 1).
TABLE 1. Mean percentage (±SEM) of cell nucleus covered by silver grains in different brain regions of intact luteal ewes (Intact, n = 7), OVX ewes (OVX, n = 8), OVX ewes bearing progesterone implants (OVX + P, n = 8)
Discussion
These studies demonstrate that progesterone treatment to OVX ewes increases dynorphin concentrations in CSF collected from the IIIV and PPD mRNA in subpopulations of dynorphin neurons. The results are thus consistent with the hypothesis that dynorphin mediates progesterone negative feedback and point to specific groups of dynorphin neurons that may contribute to the inhibition of GnRH and LH pulse frequency by this steroid.
Although progesterone treatment increased CSF dynorphin concentrations above those in OVX ewes, this effect of progesterone was not evident in luteal-phase animals. There are three possible explanations for this apparent discrepancy. First, because site of collection within the IIIV can affect estimates of peptide concentrations (32), differences in cannula placement may have increased variability among ewes and decreased the possibility of detecting significant differences. Second, it is clear that OVX does not produce a dramatic decrease in CSF dynorphin concentrations (Figs. 3 and 4), suggesting that dynorphin neurons not involved in progesterone negative feedback contribute significantly to this pool in CSF. This is not surprising, given the wide range of physiological systems influenced by hypothalamic dynorphin neurons [i.e. feeding (33), osmolality (34), lactation (35), and stress (36, 37)], but it complicates interpretation of CSF dynorphin concentrations. Third, although mean progesterone concentrations at the time of CSF collection were not different in OVX+P and luteal-phase ewes, the latter had been exposed to lower progesterone levels over the previous week because progesterone concentrations increase slowly over the first week of the luteal phase (1). Thus, an effect of progesterone may have been more evident in the OVX+P ewes, which were exposed to midluteal phase progesterone concentrations for 6–7 d. The increasing concentrations of both progesterone and CSF dynorphin over the 5-h collection period in ovary-intact ewes supports this possibility and is consistent with stimulation of dynorphin by physiological increases in progesterone concentrations.
Using in situ hybridization, we also demonstrate here that a subset of PPD mRNA-expressing cells in the ovine POA and hypothalamus, specifically those cells in the POA, AHA, and ARC, are responsive to ovarian steroids. Previous work from our laboratory has shown that over 90% of the dynorphin immunoreactive cells in these same three nuclear areas colocalize PR (15), and the same neurons are likely to contain estrogen receptor (38). In contrast, other dynorphin populations in the POA and hypothalamus showed no difference in PPD mRNA expression with hormone status. These dynorphin neurons also lack PR (15), so they are most likely involved in other actions of dynorphin such as those influencing water homeostasis (34) and the effects of stressors (36, 37).
Although treatments affected the number of cells expressing PPD mRNA in the POA, AHA, and ARC, there were no significant differences in the average percent of nucleus covered by grains among the three groups, suggesting that the amount of PPD expression per cell was not affected by treatment. This difference cannot be easily explained by a ceiling effect because marked differences were found among different areas (PVN vs. ARC) in the average percentage of nucleus covered by grains (Table 1). One interesting explanation is that only a subset of PPD-expressing cells in the POA, AHA, and ARC are responsive to progesterone; so that in the absence of progesterone, they lack adequate expression levels to be detected with our methods. After progesterone treatment, the PPD expression level increases in these cells, to match non-progesterone responsive cell levels (i.e. cells expressing PPD at high level in the absence of progesterone). This possibility could be tested by dual in situ hybridization for PPD and PR, to compare cellular PPD expression levels in response to progesterone treatment between dynorphin cells with and without PR expression.
Ovariectomy decreased the number of PPD mRNAexpressing cells in the POA, AHA, and ARC; and progesterone replacement prevented this decrease in the POA and AHA but not in the ARC (Fig. 10). One can infer from these data that: 1) progesterone actions in the POA and AHA probably account for stimulation of CSF dynorphin concentrations by progesterone (Fig. 4); and 2) another ovarian hormone, most likely estradiol (39, 40), is required to maintain PPD mRNA expression in the ARC. These regional differences in response to progesterone in OVX ewes may be explained by the observation that OVX decreased expression of PR in the ARC but not in the POA and AHA of sheep (41). They also raise the possibility that dynorphin neurons in the POA-AHA are involved in progesterone inhibition of GnRH pulse frequency in the absence of estradiol, whereas dynorphin neurons in the ARC are responsible for the enhancement of progesterone negative feedback by estradiol. It should be noted that, in assessing the physiological relevance of these different dynorphin neurons, in normal ewes progesterone always acts in the presence of estradiol (1), so that all three systems may be functional in the luteal phase.
In summary, we found that progesterone treatment alone was able to increase dynorphin concentrations in CSF and PPD mRNA expression in the POA and AHA of OVX ewes. In the ARC, there was a similar reduction in the number of cells expressing PPD in OVX ewes, but this number did not increase with progesterone treatment. These correlative data, in conjunction with earlier functional work (i.e. the stimulatory effects of a -antagonist during the luteal phase), support the hypothesis that dynorphin neurons mediate progesterone negative feedback and point to different populations of dynorphin neurons in the AHA-POA and ARC as playing important and possibly distinct roles in this action of progesterone.
Acknowledgments
We thank Drs. Gordon Niswender and Al Parlow and the National Pituitary Agency for reagents used in RIA, and Sarah Beemer and Kerie Miller for animal care.
References
Goodman R 1994 Neuroendocrine control of the ovine estrous cycle. In: Knobil ENJ, ed. The physiology of reproduction. New York: Raven Press; 659–710
Hotchkiss J, Knobil E 1994 The menstrual cycle and its neuroendocrine control. In: Knobil E, Neill J, eds. The physiology of reproduction. New York: Raven Press; 711–750
Moenter SM, Caraty A, Locatelli A, Karsch FJ 1991 Pattern of gonadotropin-releasing hormone (GnRH) secretion leading up to ovulation in the ewe: existence of a preovulatory GnRH surge. Endocrinology 129:1175–1182
Filicori M, Flamigni C, Campaniello E, Ferrari P, Meriggiola MC, Michelacci L, Pareschi A, Valdiserri A 1989 Evidence for a specific role of GnRH pulse frequency in the control of the human menstrual cycle. Am J Physiol 257:E930–E936
Skinner DC, Evans NP, Delaleu B, Goodman RL, Bouchard P, Caraty A 1998 The negative feedback actions of progesterone on gonadotropin-releasing hormone secretion are transduced by the classical progesterone receptor. Proc Natl Acad Sci USA 95:10978–10983
King JC, Tai DW, Hanna IK, Pfeiffer A, Haas P, Ronsheim PM, Mitchell SC, Turcotte JC, Blaustein JD 1995 A subgroup of LHRH neurons in guinea pigs with progestin receptors is centrally positioned within the total population of LHRH neurons. Neuroendocrinology 61:265–275
Skinner DC, Caraty A, Allingham R 2001 Unmasking the progesterone receptor in the preoptic area and hypothalamus of the ewe: no colocalization with gonadotropin-releasing neurons. Endocrinology 142:573–579
Sterling RJ, Gasc JM, Sharp PJ, Tuohimaa P, Baulieu EE 1984 Absence of nuclear progesterone receptor in LH releasing hormone in laying hens. J Endocrinol 102:R5–R7
Warembourg M, Leroy D, Peytevin J, Martinet L 1998 Estrogen receptor and progesterone receptor-immunoreactive cells are not co-localized with gonadotropin-releasing hormone in the brain of the female mink (Mustela vison). Cell Tissue Res 291:33–41
Yang K, Haynes NB, Lamming GE, Brooks AN 1988 Ovarian steroid hormone involvement in endogenous opioid modulation of LH secretion in mature ewes during the breeding and non-breeding seasons. J Reprod Fertil 83:129–139
Kalra SP 1993 Mandatory neuropeptide-steroid signaling for the preovulatory luteinizing hormone-releasing hormone discharge. Endocr Rev 14:507–538
Ferin M, Vande Wiele R 1984 Endogenous opioid peptides and the control of the menstrual cycle. Eur J Obstet Gynecol Reprod Biol 18:365–373
Gindoff PR, Jewelewicz R, Hembree W, Wardlaw S, Ferin M 1988 Sustained effects of opioid antagonism during the normal human luteal phase. J Clin Endocrinol Metab 66:1000–1004
Goodman RL, Coolen LM, Anderson GM, Hard SL, Valent MV, Connors JM, Fitzgerald ME, Lehman MM 2004 Evidence that dynorphin plays a major role in mediating progesterone negative feedback on gonadotropin-releasing hormone neurons in sheep. Endocrinology 145:2959–2967
Foradori CD, Coolen LM, Fitzgerald ME, Skinner DC, Goodman RL, Lehman MN 2002 Colocalization of progesterone receptors in parvicellular dynorphin neurons of the ovine preoptic area and hypothalamus. Endocrinology 143:4366–4374
Silverman AJ, Livne I, Witkin JW 1994 The gonadotropin-releasing hormone (GnRH) neuronal systems: immunocytochemistry and in situ hybridization. In: Knobil E, Neill JD, eds. The physiology of reproduction. 2nd ed. Vol 1. New York: Raven Press; 1683–1709
Skinner DC, Caraty A, Malpaux B, Evans NP 1997 Simultaneous measurement of gonadotropin-releasing hormone in the third ventricular cerebrospinal fluid and hypophyseal portal blood of the ewe. Endocrinology 138:4699–4704
Whisnant CS, Curto K, Goodman RL 1992 Immunocytochemical localization of ?-endorphin and gonadal steroid regulation of proopiomelanocortin messenger ribonucleic acid in the ewe. Neuroendocrinology 56:812–821
Broad KD, Kendrick KM, Sirinathsingh DJS, Keverne EB 1993 Changes in Pro-opiomelanocortin and pre-proenkephalin mRNA levels in the ovine brain during pregnancy, parturition and lactation and in response to oestrogen and progesterone. J Neuroendocrinol 5:711–719
Treiser SL, Wardlaw SL 1992 Estradiol regulation of proopiomelanocortin gene expression and peptide content in the hypothalamus. Neuroendocrinology 55:167–173
Wilcox JN, Roberts JL 1985 Estrogen decreases rat hypothalamic proopiomelanocortin messenger ribonucleic acid levels. Endocrinology 117:2392–2396
Wise PM, Scarbrough K, Weiland NG, Larson GH 1990 Diurnal pattern of proopiomelanocortin gene expression in the arcuate nucleus of proestrous, ovariectomized, and steroid-treated rats: a possible role in cyclic luteinizing hormone secretion. Mol Endocrinol 4:886–892
Borsook D, Hyman SE 1995 Proenkephalin gene regulation in the neuroendocrine hypothalamus: a model of gene regulation in the CNS. Am J Physiol 269:E393–E408
Goodman RL 1978 A quantitative analysis of the physiological role of estradiol and progesterone in control of tonic and surge secretion of luteinizing hormone in the rat. Endocrinology 102:142–150
Karsch FJ, Legan SJ, Hauger RL, Foster DL 1977 Negative feedback action of progesterone on tonic luteinizing hormone secretion in the ewe: dependence on the ovaries. Endocrinology 101:800–806
Deaver DR, Stilley NJ, Dailey RA, Inskeep EK, Lewis PE 1986 Concentrations of ovarian and pituitary hormones following prostaglandin F2-induced luteal regression in ewes varies with day of the estrous cycle at treatment. J Anim Sci 62:422–427
Whisnant CS, Goodman RL 1988 Effects of an opioid antagonist on pulsatile luteinizing hormone secretion in the ewe vary with changes in steroid negative feedback. Biol Reprod 39:1032–1038
Anderson GM, Connors JM, Hardy SL, Valent M, Goodman RL 2001 Oestradiol microimplants in the ventromedial preoptic area inhibit secretion of luteinising hormone via dopaminergic neurons in anoestrous ewes. J Neuroendocrinol 13:1051–1058
Skinner DC, Malpaux B, Delaleu B, Caraty A 1995 Luteinizing hormone (LH)-releasing hormone in third ventricular cerebrospinal fluid of the ewe: correlation with LH pulses and the LH surge. Endocrinology 136:3230–3237
Foradori CD, Goodman RL, Lehman MN, Distribution of preprodynorphin mRNA and dynorphin-A immunoreactivity in the sheep preoptic area and hypothalamus. Neuroscience, in press
Goodman RL, Karsch FJ 1980 Pulsatile secretion of luteinizing hormone: differential suppression by ovarian steroids. Endocrinology 107:1286–1290
Skinner DC, Caraty A 2002 Measurement and possible function of GnRH in cerebrospinal fluid in ewes. Reprod Suppl 59:25–39
Baile CA, McLaughlin CL, Buonomo FC, Lauterio TJ, Marson L, Della-Fera MA 1987 Opioid peptides and the control of feeding in sheep. Fed Proc 46:173–177
Shimizu H, Shimomura Y, Negishi M, Kobayashi I, Kobayashi S 1989 Dynorphin A (1–13), microinjected into the preoptic area, stimulates water intake in rats. Life Sci 45:25–30
Kim EM, Kotz CM, Welch CC, Grace MK, Billington CJ, Levine AS 1997 Lactation decreases mRNA levels of opioid peptides in the arcuate nucleus of the rat. Brain Res 769:303–308
Suh HW, Song DK, Huh SO, Kim YH 2000 Involvement of dynorphin in immobilization stress-induced antinociception in the mouse. Eur Neuropsychopharmacol 10:407–413
Matthews SG, Parrott RF, Sirinathsinghji DJ 1993 Isolation- and dehydration-induced changes in neuropeptide gene expression in the sheep hypothalamus. J Mol Endocrinol 11:181–189
Dufourny L, Skinner DC 2003 Progesterone receptor, estrogen receptor , and the type II glucocorticoid receptor are coexpressed in the same neurons of the ovine preoptic area and arcuate nucleus: a triple immunolabelling study. Biol Reprod 67:1605–1612
Karsch FJ, Legan SJ, Ryan KD, Foster DL 1980 Importance of estradiol and progesterone in regulating LH secretion and estrous behavior during the sheep estrous cycle. Biol Reprod 23:404–413
Goodman RL, Bittman EL, Foster DL, Karsch FJ 1981 The endocrine basis of the synergistic suppression of luteinizing hormone by estradiol and progesterone. Endocrinology 109:1414–1417
Scott CJ, Pereira AM, Tilbrook AJ, Rawson JA, Clarke IJ 2001 Changes in preoptic and hypothalamic levels of progesterone receptor mRNA across the oestrous cycle of the ewe. J Neuroendocrinol 13:401–406
Karsch FJ, Cummins JT, Thomas GB, Clarke IJ 1987 Steroid feedback inhibition of pulsatile secretion of gonadotropin-releasing hormone in the ewe. Biol Reprod 36:1207–1218
Evans NP, Dahl GE, Glover BH, Karsch FJ 1994 Central regulation of pulsatile gonadotropin-releasing hormone (GnRH) secretion by estradiol during the period leading up to the preovulatory GnRH surge in the ewe. Endocrinology 134:1806–1811
Goodman RL, Parfitt DB, Evans NP, Dahl GE, Karsch FJ 1995 Endogenous opioid peptides control the amplitude and shape of gonadotropin-releasing hormone pulses in the ewe. Endocrinology 136:2412–2420
Goodman RL, Gibson M, Skinner DC, Lehman MN 2002 Neuroendocrine control of pulsatile GnRH secretion during the ovarian cycle: evidence from the ewe. Reprod Suppl 59:41–56
Chavkin C, James IF, Goldstein A 1982 Dynorphin is a specific endogenous ligand of the kappa receptor. Science 215:413–415(Chad D. Foradori, Robert )
Address all correspondence and requests for reprints to: Dr. Michael N. Lehman, Department of Cell Biology, Neurobiology, and Anatomy, University of Cincinnati College of Medicine, Cincinnati, Ohio 45267-0521. E-mail: michael.lehman@uc.edu.
Abstract
Recent studies suggest that the endogenous opioid peptide, dynorphin, is an important mediator of progesterone negative feedback on GnRH pulse frequency in the ewe. These experiments tested this hypothesis by examining the effects of progesterone on dynorphin A concentrations in cerebrospinal fluid (CSF) collected from the third ventricle and expression of preprodynorphin (PPD) mRNA in hypothalamic nuclei. CSF was collected every 10 min for 5 h in three groups of ewes: 1) ovary-intact ewes during the luteal phase (d 6–7 of estrous cycle); 2) ewes 6–7 d after ovariectomy (OVX); and 3) OVX ewes treated for 6–7 d with implants that produced luteal-phase progesterone levels. Diencephalic tissue from these ewes was then collected and processed for in situ hybridization using an ovine cDNA probe against PPD. Progesterone treatment increased dynorphin A concentrations in CSF over that observed in untreated OVX ewes; CSF dynorphin A concentrations in ovary-intact ewes were midway between the other groups. OVX significantly decreased the number of PPD mRNA-expressing cells in the preoptic area (POA), anterior hypothalamic area (AHA), and arcuate nucleus (ARC), with no change seen in any other PPD-expressing nuclei. Progesterone treatment of OVX ewes restored PPD expression in the POA and AHA to levels seen in luteal-phase animals but had no effect on PPD expression in the ARC. These results are consistent with the hypothesis that progesterone acts via dynorphin neurons to inhibit pulsatile GnRH secretion and point to dynorphin neurons in the POA, AHA, and ARC as potential mediators of this action during the luteal phase.
Introduction
THE SECRETION OF GnRH into the pituitary portal blood supply is the final common pathway responsible for neuroendocrine control of reproduction. The activity of GnRH neurons, in turn, is controlled by the positive and negative feedback actions of estradiol and progesterone (1, 2, 3, 4). Specifically, elevated estradiol triggers the preovulatory GnRH surge at the end of the follicular phase, while estradiol and progesterone act singly, and in concert, to inhibit the episodic secretion of GnRH that occurs throughout the rest of the cycle.
Although both estradiol and progesterone inhibit GnRH pulses, they do so by different mechanisms. Estradiol alone inhibits LH (see Ref. 31) and GnRH (see Ref. 43) pulse amplitude, whereas progesterone by itself inhibits LH (see Ref. 31) and GnRH (5) pulse frequency, an action that is enhanced by estradiol (see Ref. 40). In sheep, this action of progesterone appears to be mediated by the classical nuclear progesterone receptor (PR) (5), but PR are generally not found in GnRH neurons in this and other species (6, 7, 8, 9). Thus, other systems most likely transmit the actions of progesterone to GnRH neurons. There is now general agreement that endogenous opioid peptides (EOP) are one of the major systems that mediate progesterone negative feedback in sheep (1, 10), rats (11), and primates (2, 12, 13). In contrast, there is clear evidence that EOP do not mediate the inhibition of GnRH pulse amplitude by estradiol in sheep (see Ref. 44), and there is some evidence implicating noradrenergic neurons in this action of estradiol (see Ref. 45).
Recent work has raised the possibility that dynorphin A1–17 may be the primary EOP involved in progesterone negative feedback. Dynorphin A1–17 is part of a family of EOP derived from preprodynorphin (PPD) that also includes dynorphin A1–8 and dynorphin B. Specifically, three lines of evidence have implicated dynorphin A, which is the endogenous ligand for the EOP receptor (see Ref. 46): 1) almost all GnRH neurons in the medial basal hypothalamus, and about 50% of those elsewhere in the sheep brain, receive synaptic inputs from dynorphin A-containing terminals (14); 2) an EOP receptor antagonist consistently increases LH pulse frequency in luteal phase ewes (14); and 3) over 90% of the dynorphin A-positive perikarya in the preoptic area (POA), anterior hypothalamus (AHA), and arcuate nucleus (ARC) contain PR (15).
If this working hypothesis is correct, then progesterone should increase dynorphin release from the synapses that terminate on GnRH neurons. However, this prediction is difficult to test directly because of the dispersed nature of GnRH perikarya (16). Therefore, we have chosen two less-direct approaches to testing it. First, we assessed the effect of progesterone on dynorphin A concentrations in the cerebrospinal fluid (CSF) collected from the third ventricle (IIIV); third ventricular CSF GnRH levels in the sheep correlate well with its endogenous secretion rates (17). Second, we tested whether progesterone would increase PPD mRNA levels in distinct subpopulations of dynorphin-containing neurons, an approach that has been useful in assessing the effects of steroids on other EOP in the sheep (18, 19) and rat (20, 21, 22, 23).
Although there are similarities in the EOP mediation of progesterone negative feedback in sheep, rats, and primates, there is one difference in this action of progesterone among these species. In primates (2) and rats (24), the negative feedback action of progesterone requires the presence of estradiol; whereas in sheep, progesterone alone inhibits tonic LH secretion in acutely ovariectomized (OVX) animals (25). Consequently, many studies in sheep have examined this action of progesterone in the absence of estradiol (1, 2, 5, 10, 31); and there is evidence that, by itself, progesterone can affect the expression of the mRNA for EOP in sheep (18, 19). Because progesterone negative feedback does not require estradiol and estradiol negative feedback does not involve EOP, in this study we examined the actions of progesterone on dynorphin A concentrations in CSF and PPD mRNA levels in the absence of estradiol.
Materials and Methods
Animals
Adult Suffolk ewes were maintained in an open barn with free access to water and fed once daily with a maintenance regimen of silage supplemented with grain. This study was performed during the breeding season (January and early February) in 23 ewes that had demonstrated at least two normal 16- to 17-d estrous cycles (determined by monitoring estrous behavior with a vasectomized ram). They were moved to indoor facilities 2–3 d before experimentation. Once indoors, the animals were housed two ewes/pen under a photoperiod similar to that occurring outdoors. The routine handling and experimental procedures involving animals were approved by the West Virginia University Animal Care and Use Committee.
Experimental protocols
Ovarian cycles were synchronized by injection of prostaglandin F2, 7 d apart (26). Three days after the second prostaglandin F2 injection (approximately d 0 of next ovarian cycle), ewes were either OVX (n = 16) or sham OVX (n = 7) and a cannula inserted into the IIIV. Eight of the OVX ewes received two SILASTIC-brand (Dow Corning Corp., Midland, MI) packets of progesterone placed sc just after OVX to produce midluteal phase P concentrations (27). Blood and CSF samples were then collected 6–7 d after surgery. There were thus three treatment groups: ovary-intact (n = 7) on d 6 or 7 of the estrous cycle, OVX (n = 8), and OVX+P (n = 8).
Surgeries
Ovariectomies were preformed by midventral laparotomy using sterile procedures under gas anesthesia (oxygen+nitrous oxide, supplemented with halothane as needed). After OVX (or sham OVX, which consisted of midventral incision), a 16-gauge stainless steel cannula was placed into the IIIV using a stereotaxic procedure as previously described in detail (28). Briefly, the surface of the skull was exposed, a 1-cm-diameter hole drilled, and the sagittal sinus ligated and cut. Radio-opaque dye was then injected into one lateral ventricle, and frontal and lateral x-rays were used to guide the placement of the cannula with its tip about 5 mm dorsal to the median eminence. Placement in the IIIV was confirmed by flow, or withdrawal of CSF, from the cannula. The cannula was then cemented to the skull with screws and dental acrylic, plugged, and covered with a protective cap. All ewes were monitored closely post operation, and only those that showed complete recovery (based on temperature, food intake, and level of arousal) within 3 d were used in these experiments.
Blood and CSF collections
Collection of CSF was based on the procedure described by Skinner et al. (29). The day before collection, one jugular vein was catheterized and a panel used to restrict each animal’s movement so that it could lie down but not turn around. The next morning, polyethylene tubing (0.38-mm inside diameter, 1.09-mm outside diameter; VWR International, Bridgeport NJ) was inserted down the cannula so that it extended about 1 mm beyond its end into the ventricle. This tubing was connected via silicone rubber tubing (1.0-mm inside diameter, 2.0-mm outside diameter; Sil-Med Corp., Taunton, MA) to a Gilson peristaltic pump, and samples were collected every 10 min for 5 h. Approximately 0.5 ml CSF was collected on ice continuously over the 10-min period and was immediately frozen after collection. Heparinized blood samples (5 ml) were taken every 10 min, from the catheter previously placed into the jugular vein, for 5 h. In a few cases, flow through the peristalic pump failed and samples were continued to be collected directly from the cannula with a syringe every 10 min. In six ewes, only a few CSF samples could be collected; these data were not included in the analyses, leaving five to six animals/group. Plasma was stored at –20 C and CSF samples stored at –80 C until assayed.
Tissue collection
The day after CSF and blood sample collection, fresh tissue was collected for in situ hybridization analysis of PPD mRNA expression. Animals were given an anesthetic dose of sodium pentobarbital (1 g, iv), and the protective cap and cannula were removed to facilitate rapid collection of tissue. The ewes were then given a lethal dose of pentobarbital (2 g, iv) and decapitated; and a tissue block containing the septal region, POA, and hypothalamus was rapidly dissected out and placed on dry ice. After freezing, the blocks were shipped on dry ice to the University of Cincinnati and stored at –80 C until sectioned. Coronal sections (20 μm) were cut on a cryostat, thaw mounted on SuperFrost plus slides (Fisher Scientific, Pittsburgh, PA), and stored at –80 C until processed for in situ hybridization.
RIAs
LH was measured in duplicate aliquots of all plasma samples using a previously described RIA (27), except that we iodinated-purified ovine LH provided by Dr. Parlow and the National Hormone and Peptide Program for tracer. Assay sensitivity averaged 0.18 ng/ml (NIH-S24). Intraassay and interassay coefficients of variation (CVs) averaged 9% and 13%, respectively. Progesterone was measured in 100-μl aliquots of plasma samples collected at three time points spanning the 5-h collection period using a commercially available kit (Diagnostic Systems Laboratories, Inc., Webster, TX) previously validated for use in sheep (28). All samples were measured in a single assay, with a sensitivity of 0.02 ng/ml and intraassay CV of 9%.
Dynorphin concentrations in CSF were measured using a commercial kit (Peninsula Laboratories, Inc., Belmont, CA). Briefly, 100 μl rabbit dynorphin A antisera was added to the standard curve (in triplicate, ranging from 1–32 pg) and duplicate aliquots of CSF (usually 150 μl) that had been lyophilized and reconstituted in 100 μl assay buffer. After an overnight incubation at 4 C, iodinated-dynorphin A (100 μl; 15,000 cpm) was added, and incubation at 4 C continued for another 24 h. Antibody-bound radioactivity was separated from unbound with donkey antirabbit -globulin on d 3. This assay has minimal cross-reactivity (<1%) with dynorphin A (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13) and no cross-reactivity with dynorphin B, ?-endorphin, or leu-enkephalin. The assay sensitivity averaged 1.0 ± 0.2 pg (n = 6), and the ED50 was 12.1 ± 0.6 pg (n = 6). Pools of CSF from three different ewes showed parallelism with the standard curve (Fig. 1). Almost all (>95%) unknown samples fell between 80% and 50% displacement of iodinated dynorphin, and the intraassay CVs of variation at these points on the standard curve were 19 ± 0.8% and 12.1 ± 1.1%, respectively; interassay CV averaged 12% for the six assays in which CSF levels were monitored.
FIG. 1. Displacement of iodinated-dynorphin from dynorphin-A antibody by 1–32 pg dynorphin (lower plot and x-axis) and 50, 100, and 200 μl of three pools of CSF (top axis and upper plots).
In situ hybridization
Probes were produced using a cDNA sequence of a portion of the ovine PPD previously described (30). Antisense and sense RNA fragments were generated from linearized templates (2 μg) using T7 and SP6 RNA polymerases (Promega, Madison, WI), respectively, in a transcription reaction containing: 5x transcription buffer (Promega), 5 mM ribonucleotide triphosphates (GTP, CTP, ATP), 100 μM UTP, ribonuclease (RNase) inhibitor (Rnasin, Promega, N211A), 100 mM dithiothreitol, and 100 μCi 35S-UTP (3000 Ci/mmol; NEN Life Science Products, Boston, MA) for 2 h at 37 C. Samples were then incubated with Dnase-I (Sigma, St. Louis, MO) at 37 C for 15 min to remove template. The reaction was stopped with 0.5 EDTA. Unincorporated isotopes were removed using spin columns (Roche, Indianapolis, IN). Probes were diluted in Tris, EDTA, containing 10 mg/ml Torula RNA (Sigma), denatured at 90 C for 3 min, and combined with hybridization buffer containing 50x Reinhardt’s solution, deionized formamide, 50% dextran sulfate, 5 M NaCl, 1 M Tris (pH 8.0), 0.5 EDTA, and 1 M dithiothreitol.
Just before use, slides containing tissue sections were removed from the freezer, placed in racks, and dried for 30–40 sec. Sections were then fixed for 5 min in 4% paraformaldehyde in phosphate buffer (PB, pH 7.4) at room temperature and then rinsed twice in PB, while stirring. Slides were dipped briefly in water, then in triethanolamine, and finally, soaked in triethanolamine plus acetic anhydride for 10 min, while stirring. Slides were then soaked in 2x saline sodium citrate (SSC: 0.15 M NaCl, 0.015 M sodium citrate); 70%, 95%, and 100% ethanol, respectively; followed by delipidation in chloroform for 5 min. Slides were soaked for 3 min each in 100% and 95% ethanol and then air-dried. Once dried, hybridization buffer with labeled probe (1 x 106 cpm/slide) was applied to the slides in a vol of 150 μl. Calculations based on molar amount of probe and estimates of tissue RNA indicated that this probe is more than 1000-fold in excess of endogenous levels. Slides were coverslipped with Parafilm (Fisher Scientific) and sealed with rubber cement and hybridized overnight at 46 C. On the following day, coverslips were removed, and the slides were washed for 15 min in 4x SSC. Slides were then incubated for 30 min at 37 C in RNase buffer (1 M Tris, pH 8.0; 5 M NaCl; 0.5 M EDTA) containing RNase-A (0.03 mg/ml; Sigma) followed by incubation for 30 min at 37 C in buffer without RNase-A. Next, the slides were washed at room temperature under increasingly stringent conditions (2x SSC, 0.1x SSC [59 C, twice, 30 min each], 0.1 x SSC [room temperature, twice]). Finally, slides were dehydrated for 3 min each in 50, 85, and 100% ethanol and then air-dried. Slides were then dipped in Kodak NTB3 emulsion, dried, and exposed in the dark for 21 d at 4 C. The emulsion was developed in Dektol (Kodak, Rochester, NY). The sections were counterstained with cresyl violet, dehydrated, and coverslipped with DPX (EM Sciences, Fort Washington, PA). Sense controls and RNase-pretreated sections showed only nonspecific labeling and background levels comparable with antisense-treated sections.
Data analysis
Number of cells expressing PPD.
Sections were coded so that analysis would be preformed blind to the condition of the animals. The locations of PPD-expressing cells were identified in the POA and hypothalamus of each ewe under dark-field and bright-field microscopy using a Leica microscope. Every tenth section was analyzed. Regions chosen for analysis were those shown to contain large numbers of PPD mRNAexpressing cells in our previous study (30): the POA, AHA, bed nucleus of the stria terminalis (BNST), supraoptic nucleus (SON), paraventricular nucleus (PVN), ventromedial nucleus (VMN), dorsomedial nucleus of the hypothalamus (DMH), and ARC. Three sections were analyzed for each area. Individual circular deposits of silver grains were considered positive for PPD if grains were 5x over background density. Images of labeled material were captured using a digital camera (Magnafire; Optronics, Goleta, CA) attached to a Leica microscope (Deerfield, IL) and imported into Adobe Photoshop 7.0 (Adobe Systems, San Jose, CA). Images were not altered in any way except for minor adjustments of brightness and contrast. Low-power drawings of the sections analyzed were made with a camera lucida to help provide orientation for the dark-field images.
Percent of nucleus covered by grains.
For every area analyzed, 12 cells with clear, Nissl-stained nuclei were randomly chosen, and high-powered (x40) bright-field photomicrographs were taken of each (Fig. 2). Because Nissl staining of the cytosol was uneven, it was impossible to determine whether grains overlying regions outside the stained nuclei were due to the cell in question or neighboring cells. Thus, we chose a more conservative quantitation method and analyzed silver grains over the nucleus. Using NIH Image, the area of the Nissl-stained nucleus of each cell was outlined (e.g. Fig. 2), and the total area of the nucleus was found. A standard threshold was chosen so that no pixels were attributed to the Nissl-stained nucleus and only pixels associated with silver grains were measured. The total number of silver grain pixels within the area of the nucleus was determined. Then the same shape traced around the nucleus was used to analyze an area lacking Nissl staining, and the total number of pixels was measured within this area. This created a background pixel number. The total number of pixels attributed to background silver grain deposits was subtracted from the number of pixels of the silver grains over the nucleus. This number was then divided by the total area outlined by the nucleus to arrive at the percent of the nucleus covered by grains. The percent grain coverage for the 12 random cells was averaged to obtain a mean percentage of the nucleus covered by grains for each area in each animal.
FIG. 2. High-power bright-field photomicrograph showing an example of a PPD-positive cell in the ARC. Silver grains are seen overlying the blue, Nissl-stained nucleus of this cell (outlined area) as well as surrounding it. Scale bar, 10 μm.
Statistical analysis
Hormones and CSF dynorphin concentrations.
LH pulses were identified using previously published criteria (31) and differences among treatment groups in LH pulse frequency evaluated by ANOVA. Mean differences in dynorphin concentrations in CSF and P in serum, the number of cells expressing PPD, and the average percent of nucleus covered by grains among treatment groups were all analyzed by one-way ANOVA followed by Tukey’s post hoc test. Changes in serum progesterone and CSF dynorphin concentrations over the 5-h period of collection, within each treatment group, were analyzed by one-way ANOVA with repeated measure and linear regression analysis, respectively. P < 0.05 was considered statistically significant for all analyses.
Results
Serum hormone and CSF dynorphin concentrations
As expected, there was no difference in the peripheral progesterone concentrations between the intact and OVX+P groups, whereas the OVX ewes showed levels of progesterone at the lower limit of detection (Fig. 3). LH pulse frequency in the ovary-intact and OVX+P groups averaged approximately 1 pulse/5 h and was significantly increased to 4 pulses/5 h in the OVX group (Figs. 3 and 4).
FIG. 3. Histogram comparing the concentration of CSF dynorphin, serum levels of progesterone, and LH pulse frequency (FREQ) in intact luteal controls (Intact, n = 6), OVX (n = 5), and OVX ewes bearing progesterone implants (OVX+P, n = 6). Different letters signify P < 0.05.
FIG. 4. Concentrations of plasma LH (bottom panels) and of dynorphin in CSF collected from the IIIV (top panels) of an ovary-intact ewe (left panels), an OVX (middle panels) ewe, and an OVX ewe treated with luteal-phase P levels (right panels). Solid circles in plots of LH indicate peaks of LH pulses.
Dynorphin A concentrations in CSF collected from the IIIV showed marked temporal variations, some as great as 50–100% over a 10- to 20-min sampling period, suggestive of an episodic mode of release (Fig. 4). However, there was no obvious correlation between the patterns of dynorphin in CSF and LH pulses in peripheral circulation or progesterone treatment (Fig. 4). The OVX ewes treated with progesterone had a significantly higher mean dynorphin concentration in CSF compared with that in OVX animals (Fig. 3). Intact ewes had intermediate CSF dynorphin levels and did not differ significantly from levels in either the OVX or OVX+P groups.
In OVX and OVX+P animals, peripheral progesterone concentrations were relatively constant during the 5-h collection period; but in the ovary-intact ewes, progesterone levels significantly increased from 1.4 ± 0.2 to 2.2 ± 0.3 ng/ml over this period (Fig. 5). During this same period, there was a gradual increase in CSF dynorphin concentrations in ovary-intact ewes (Fig. 5) that was statistically significant (P < 0.001) by linear regression analysis. There were no significant trends over the 5-h collection period in CSF dynorphin concentrations in the OVX or OVX+P groups (data not shown).
FIG. 5. Mean (±SEM) serum progesterone (PROG) concentrations (bars) and dynorphin concentrations (solid circles) in CSF samples collected from ovary-intact ewes during the luteal phase of the estrous cycle (n = 6).
Expression of PPD mRNA
As previously described (30), PPD mRNA-expressing cells were found in a number of nuclei of the ovine POA and medial basal hypothalamus, including the POA, AHA, BNST, SON, PVN, VMN, DMH, and ARC. Three of these areas, the POA (Fig. 6, A and B), AHA (Fig. 6, D–F), and ARC (Fig. 7), showed a significant change in the number of cells expressing PPD mRNA among the three experimental groups (see Fig. 10). Other areas, including the SON (Fig. 8), PVN (Fig. 9), VMN, DMH, and BNST, showed no significant differences among groups in the number of cells expressing PPD mRNA (Fig. 10).
FIG. 6. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the ventromedial POA (A–C) and AHA (D–E) of intact luteal ewes (A and D), OVX ewes (B and E), and OVX ewes bearing implants that maintain luteal phase levels of progesterone (OVX+P) (C and F). For orientation, camera lucida drawings of the sections from the intact ewes (A and D) are displayed at left, and the solid boxes within these drawings show the areas depicted in the photomicrographs. Note that the number of labeled cells in the rostral portion of the PVN (rPVN) (arrows in D and E) does not change. OVLT, Organum vasculosum of the lamina terminalis; 3v, IIIV; ac, anterior commissure; oc, optic chiasm. Scale bar, 200 μm.
FIG. 7. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the ARC of intact luteal phase (A), OVX (B), and OVX+P (C) ewes. The boxed area in the camera lucida drawing at left shows the region depicted in the photomicrographs. fx, Fornix; mt, mammillothalamic tract. Scale bar, 200 μm.
FIG. 10. Histogram comparing the mean (±SEM) number of PPD mRNA-expressing cells in the POA, AHA, ARC, PVN, VMN, DMH of luteal phase (Intact, n = 7), OVX (n = 8), and OVX ewes bearing progesterone implants (+P, n = 8). Asterisks, P < 0.05.
FIG. 8. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the SON (A–C) of intact luteal controls (A), OVX ewes (B), and OVX+P ewes (C). The boxed area in the camera lucida drawing at left shows the region depicted in the photomicrographs. Scale bar, 200 μm.
FIG. 9. Low-power dark-field photomicrographs showing PPD mRNA-expressing cells in the PVN (A–C) of intact luteal controls (A), OVX ewes (B), and OVX+P ewes (C). The boxed area in the camera lucida drawing at left shows the region depicted in the photomicrographs. ot, Optic tract. Scale bar, 200 μm.
Compared with intact animals, OVX ewes showed almost a 50% decrease in the number of PPD mRNA-expressing cells in the POA and AHA and a 30% decrease in the ARC (Figs. 6, A–C, and 10). Progesterone treatment of OVX ewes increased the number of PPD mRNA-expressing cells in the POA and AHA to levels not significantly different from intact controls (Figs. 6 and 10). However, in the ARC, the number of cells expressing PPD mRNA in the P-treated group did not return to the intact level, and remained at the OVX level of expression (Figs. 7, A–C, and 10). Neither OVX nor progesterone treatment of OVX ewes affected the percentage of the cell nucleus covered by silver grains for any of the areas analyzed (Table 1).
TABLE 1. Mean percentage (±SEM) of cell nucleus covered by silver grains in different brain regions of intact luteal ewes (Intact, n = 7), OVX ewes (OVX, n = 8), OVX ewes bearing progesterone implants (OVX + P, n = 8)
Discussion
These studies demonstrate that progesterone treatment to OVX ewes increases dynorphin concentrations in CSF collected from the IIIV and PPD mRNA in subpopulations of dynorphin neurons. The results are thus consistent with the hypothesis that dynorphin mediates progesterone negative feedback and point to specific groups of dynorphin neurons that may contribute to the inhibition of GnRH and LH pulse frequency by this steroid.
Although progesterone treatment increased CSF dynorphin concentrations above those in OVX ewes, this effect of progesterone was not evident in luteal-phase animals. There are three possible explanations for this apparent discrepancy. First, because site of collection within the IIIV can affect estimates of peptide concentrations (32), differences in cannula placement may have increased variability among ewes and decreased the possibility of detecting significant differences. Second, it is clear that OVX does not produce a dramatic decrease in CSF dynorphin concentrations (Figs. 3 and 4), suggesting that dynorphin neurons not involved in progesterone negative feedback contribute significantly to this pool in CSF. This is not surprising, given the wide range of physiological systems influenced by hypothalamic dynorphin neurons [i.e. feeding (33), osmolality (34), lactation (35), and stress (36, 37)], but it complicates interpretation of CSF dynorphin concentrations. Third, although mean progesterone concentrations at the time of CSF collection were not different in OVX+P and luteal-phase ewes, the latter had been exposed to lower progesterone levels over the previous week because progesterone concentrations increase slowly over the first week of the luteal phase (1). Thus, an effect of progesterone may have been more evident in the OVX+P ewes, which were exposed to midluteal phase progesterone concentrations for 6–7 d. The increasing concentrations of both progesterone and CSF dynorphin over the 5-h collection period in ovary-intact ewes supports this possibility and is consistent with stimulation of dynorphin by physiological increases in progesterone concentrations.
Using in situ hybridization, we also demonstrate here that a subset of PPD mRNA-expressing cells in the ovine POA and hypothalamus, specifically those cells in the POA, AHA, and ARC, are responsive to ovarian steroids. Previous work from our laboratory has shown that over 90% of the dynorphin immunoreactive cells in these same three nuclear areas colocalize PR (15), and the same neurons are likely to contain estrogen receptor (38). In contrast, other dynorphin populations in the POA and hypothalamus showed no difference in PPD mRNA expression with hormone status. These dynorphin neurons also lack PR (15), so they are most likely involved in other actions of dynorphin such as those influencing water homeostasis (34) and the effects of stressors (36, 37).
Although treatments affected the number of cells expressing PPD mRNA in the POA, AHA, and ARC, there were no significant differences in the average percent of nucleus covered by grains among the three groups, suggesting that the amount of PPD expression per cell was not affected by treatment. This difference cannot be easily explained by a ceiling effect because marked differences were found among different areas (PVN vs. ARC) in the average percentage of nucleus covered by grains (Table 1). One interesting explanation is that only a subset of PPD-expressing cells in the POA, AHA, and ARC are responsive to progesterone; so that in the absence of progesterone, they lack adequate expression levels to be detected with our methods. After progesterone treatment, the PPD expression level increases in these cells, to match non-progesterone responsive cell levels (i.e. cells expressing PPD at high level in the absence of progesterone). This possibility could be tested by dual in situ hybridization for PPD and PR, to compare cellular PPD expression levels in response to progesterone treatment between dynorphin cells with and without PR expression.
Ovariectomy decreased the number of PPD mRNAexpressing cells in the POA, AHA, and ARC; and progesterone replacement prevented this decrease in the POA and AHA but not in the ARC (Fig. 10). One can infer from these data that: 1) progesterone actions in the POA and AHA probably account for stimulation of CSF dynorphin concentrations by progesterone (Fig. 4); and 2) another ovarian hormone, most likely estradiol (39, 40), is required to maintain PPD mRNA expression in the ARC. These regional differences in response to progesterone in OVX ewes may be explained by the observation that OVX decreased expression of PR in the ARC but not in the POA and AHA of sheep (41). They also raise the possibility that dynorphin neurons in the POA-AHA are involved in progesterone inhibition of GnRH pulse frequency in the absence of estradiol, whereas dynorphin neurons in the ARC are responsible for the enhancement of progesterone negative feedback by estradiol. It should be noted that, in assessing the physiological relevance of these different dynorphin neurons, in normal ewes progesterone always acts in the presence of estradiol (1), so that all three systems may be functional in the luteal phase.
In summary, we found that progesterone treatment alone was able to increase dynorphin concentrations in CSF and PPD mRNA expression in the POA and AHA of OVX ewes. In the ARC, there was a similar reduction in the number of cells expressing PPD in OVX ewes, but this number did not increase with progesterone treatment. These correlative data, in conjunction with earlier functional work (i.e. the stimulatory effects of a -antagonist during the luteal phase), support the hypothesis that dynorphin neurons mediate progesterone negative feedback and point to different populations of dynorphin neurons in the AHA-POA and ARC as playing important and possibly distinct roles in this action of progesterone.
Acknowledgments
We thank Drs. Gordon Niswender and Al Parlow and the National Pituitary Agency for reagents used in RIA, and Sarah Beemer and Kerie Miller for animal care.
References
Goodman R 1994 Neuroendocrine control of the ovine estrous cycle. In: Knobil ENJ, ed. The physiology of reproduction. New York: Raven Press; 659–710
Hotchkiss J, Knobil E 1994 The menstrual cycle and its neuroendocrine control. In: Knobil E, Neill J, eds. The physiology of reproduction. New York: Raven Press; 711–750
Moenter SM, Caraty A, Locatelli A, Karsch FJ 1991 Pattern of gonadotropin-releasing hormone (GnRH) secretion leading up to ovulation in the ewe: existence of a preovulatory GnRH surge. Endocrinology 129:1175–1182
Filicori M, Flamigni C, Campaniello E, Ferrari P, Meriggiola MC, Michelacci L, Pareschi A, Valdiserri A 1989 Evidence for a specific role of GnRH pulse frequency in the control of the human menstrual cycle. Am J Physiol 257:E930–E936
Skinner DC, Evans NP, Delaleu B, Goodman RL, Bouchard P, Caraty A 1998 The negative feedback actions of progesterone on gonadotropin-releasing hormone secretion are transduced by the classical progesterone receptor. Proc Natl Acad Sci USA 95:10978–10983
King JC, Tai DW, Hanna IK, Pfeiffer A, Haas P, Ronsheim PM, Mitchell SC, Turcotte JC, Blaustein JD 1995 A subgroup of LHRH neurons in guinea pigs with progestin receptors is centrally positioned within the total population of LHRH neurons. Neuroendocrinology 61:265–275
Skinner DC, Caraty A, Allingham R 2001 Unmasking the progesterone receptor in the preoptic area and hypothalamus of the ewe: no colocalization with gonadotropin-releasing neurons. Endocrinology 142:573–579
Sterling RJ, Gasc JM, Sharp PJ, Tuohimaa P, Baulieu EE 1984 Absence of nuclear progesterone receptor in LH releasing hormone in laying hens. J Endocrinol 102:R5–R7
Warembourg M, Leroy D, Peytevin J, Martinet L 1998 Estrogen receptor and progesterone receptor-immunoreactive cells are not co-localized with gonadotropin-releasing hormone in the brain of the female mink (Mustela vison). Cell Tissue Res 291:33–41
Yang K, Haynes NB, Lamming GE, Brooks AN 1988 Ovarian steroid hormone involvement in endogenous opioid modulation of LH secretion in mature ewes during the breeding and non-breeding seasons. J Reprod Fertil 83:129–139
Kalra SP 1993 Mandatory neuropeptide-steroid signaling for the preovulatory luteinizing hormone-releasing hormone discharge. Endocr Rev 14:507–538
Ferin M, Vande Wiele R 1984 Endogenous opioid peptides and the control of the menstrual cycle. Eur J Obstet Gynecol Reprod Biol 18:365–373
Gindoff PR, Jewelewicz R, Hembree W, Wardlaw S, Ferin M 1988 Sustained effects of opioid antagonism during the normal human luteal phase. J Clin Endocrinol Metab 66:1000–1004
Goodman RL, Coolen LM, Anderson GM, Hard SL, Valent MV, Connors JM, Fitzgerald ME, Lehman MM 2004 Evidence that dynorphin plays a major role in mediating progesterone negative feedback on gonadotropin-releasing hormone neurons in sheep. Endocrinology 145:2959–2967
Foradori CD, Coolen LM, Fitzgerald ME, Skinner DC, Goodman RL, Lehman MN 2002 Colocalization of progesterone receptors in parvicellular dynorphin neurons of the ovine preoptic area and hypothalamus. Endocrinology 143:4366–4374
Silverman AJ, Livne I, Witkin JW 1994 The gonadotropin-releasing hormone (GnRH) neuronal systems: immunocytochemistry and in situ hybridization. In: Knobil E, Neill JD, eds. The physiology of reproduction. 2nd ed. Vol 1. New York: Raven Press; 1683–1709
Skinner DC, Caraty A, Malpaux B, Evans NP 1997 Simultaneous measurement of gonadotropin-releasing hormone in the third ventricular cerebrospinal fluid and hypophyseal portal blood of the ewe. Endocrinology 138:4699–4704
Whisnant CS, Curto K, Goodman RL 1992 Immunocytochemical localization of ?-endorphin and gonadal steroid regulation of proopiomelanocortin messenger ribonucleic acid in the ewe. Neuroendocrinology 56:812–821
Broad KD, Kendrick KM, Sirinathsingh DJS, Keverne EB 1993 Changes in Pro-opiomelanocortin and pre-proenkephalin mRNA levels in the ovine brain during pregnancy, parturition and lactation and in response to oestrogen and progesterone. J Neuroendocrinol 5:711–719
Treiser SL, Wardlaw SL 1992 Estradiol regulation of proopiomelanocortin gene expression and peptide content in the hypothalamus. Neuroendocrinology 55:167–173
Wilcox JN, Roberts JL 1985 Estrogen decreases rat hypothalamic proopiomelanocortin messenger ribonucleic acid levels. Endocrinology 117:2392–2396
Wise PM, Scarbrough K, Weiland NG, Larson GH 1990 Diurnal pattern of proopiomelanocortin gene expression in the arcuate nucleus of proestrous, ovariectomized, and steroid-treated rats: a possible role in cyclic luteinizing hormone secretion. Mol Endocrinol 4:886–892
Borsook D, Hyman SE 1995 Proenkephalin gene regulation in the neuroendocrine hypothalamus: a model of gene regulation in the CNS. Am J Physiol 269:E393–E408
Goodman RL 1978 A quantitative analysis of the physiological role of estradiol and progesterone in control of tonic and surge secretion of luteinizing hormone in the rat. Endocrinology 102:142–150
Karsch FJ, Legan SJ, Hauger RL, Foster DL 1977 Negative feedback action of progesterone on tonic luteinizing hormone secretion in the ewe: dependence on the ovaries. Endocrinology 101:800–806
Deaver DR, Stilley NJ, Dailey RA, Inskeep EK, Lewis PE 1986 Concentrations of ovarian and pituitary hormones following prostaglandin F2-induced luteal regression in ewes varies with day of the estrous cycle at treatment. J Anim Sci 62:422–427
Whisnant CS, Goodman RL 1988 Effects of an opioid antagonist on pulsatile luteinizing hormone secretion in the ewe vary with changes in steroid negative feedback. Biol Reprod 39:1032–1038
Anderson GM, Connors JM, Hardy SL, Valent M, Goodman RL 2001 Oestradiol microimplants in the ventromedial preoptic area inhibit secretion of luteinising hormone via dopaminergic neurons in anoestrous ewes. J Neuroendocrinol 13:1051–1058
Skinner DC, Malpaux B, Delaleu B, Caraty A 1995 Luteinizing hormone (LH)-releasing hormone in third ventricular cerebrospinal fluid of the ewe: correlation with LH pulses and the LH surge. Endocrinology 136:3230–3237
Foradori CD, Goodman RL, Lehman MN, Distribution of preprodynorphin mRNA and dynorphin-A immunoreactivity in the sheep preoptic area and hypothalamus. Neuroscience, in press
Goodman RL, Karsch FJ 1980 Pulsatile secretion of luteinizing hormone: differential suppression by ovarian steroids. Endocrinology 107:1286–1290
Skinner DC, Caraty A 2002 Measurement and possible function of GnRH in cerebrospinal fluid in ewes. Reprod Suppl 59:25–39
Baile CA, McLaughlin CL, Buonomo FC, Lauterio TJ, Marson L, Della-Fera MA 1987 Opioid peptides and the control of feeding in sheep. Fed Proc 46:173–177
Shimizu H, Shimomura Y, Negishi M, Kobayashi I, Kobayashi S 1989 Dynorphin A (1–13), microinjected into the preoptic area, stimulates water intake in rats. Life Sci 45:25–30
Kim EM, Kotz CM, Welch CC, Grace MK, Billington CJ, Levine AS 1997 Lactation decreases mRNA levels of opioid peptides in the arcuate nucleus of the rat. Brain Res 769:303–308
Suh HW, Song DK, Huh SO, Kim YH 2000 Involvement of dynorphin in immobilization stress-induced antinociception in the mouse. Eur Neuropsychopharmacol 10:407–413
Matthews SG, Parrott RF, Sirinathsinghji DJ 1993 Isolation- and dehydration-induced changes in neuropeptide gene expression in the sheep hypothalamus. J Mol Endocrinol 11:181–189
Dufourny L, Skinner DC 2003 Progesterone receptor, estrogen receptor , and the type II glucocorticoid receptor are coexpressed in the same neurons of the ovine preoptic area and arcuate nucleus: a triple immunolabelling study. Biol Reprod 67:1605–1612
Karsch FJ, Legan SJ, Ryan KD, Foster DL 1980 Importance of estradiol and progesterone in regulating LH secretion and estrous behavior during the sheep estrous cycle. Biol Reprod 23:404–413
Goodman RL, Bittman EL, Foster DL, Karsch FJ 1981 The endocrine basis of the synergistic suppression of luteinizing hormone by estradiol and progesterone. Endocrinology 109:1414–1417
Scott CJ, Pereira AM, Tilbrook AJ, Rawson JA, Clarke IJ 2001 Changes in preoptic and hypothalamic levels of progesterone receptor mRNA across the oestrous cycle of the ewe. J Neuroendocrinol 13:401–406
Karsch FJ, Cummins JT, Thomas GB, Clarke IJ 1987 Steroid feedback inhibition of pulsatile secretion of gonadotropin-releasing hormone in the ewe. Biol Reprod 36:1207–1218
Evans NP, Dahl GE, Glover BH, Karsch FJ 1994 Central regulation of pulsatile gonadotropin-releasing hormone (GnRH) secretion by estradiol during the period leading up to the preovulatory GnRH surge in the ewe. Endocrinology 134:1806–1811
Goodman RL, Parfitt DB, Evans NP, Dahl GE, Karsch FJ 1995 Endogenous opioid peptides control the amplitude and shape of gonadotropin-releasing hormone pulses in the ewe. Endocrinology 136:2412–2420
Goodman RL, Gibson M, Skinner DC, Lehman MN 2002 Neuroendocrine control of pulsatile GnRH secretion during the ovarian cycle: evidence from the ewe. Reprod Suppl 59:41–56
Chavkin C, James IF, Goldstein A 1982 Dynorphin is a specific endogenous ligand of the kappa receptor. Science 215:413–415(Chad D. Foradori, Robert )