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Endocrinology Vol. 139, No. 1 57-64
Copyright © 1998 by The Endocrine Society

ARTICLES

Gonadotropin-Releasing Hormone Messenger Ribonucleic Acid Expression Changes before the Onset of the Estradiol-Induced Luteinizing Hormone Surge in the Ewe1

Thomas G. Harris2, Jane E. Robinson, Neil P. Evans, Donal C. Skinner3 and Allan E. Herbison4

Laboratory of Neuroendocrinology, The Babraham Institute, Babraham, Cambridge, United Kingdom CB2 4AT

Address all correspondence and requests for reprints to: Dr. J. E. Robinson, Department of Neurobiology, The Babraham Institute, Babraham, Cambridge, United Kingdom CB2 4AT. E-mail: jane.robinson{at}bbsrc.ac.uk


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The preovulatory LH surge in the ewe is stimulated by the massive and sustained release of GnRH into the pituitary portal vessels. This study has examined the temporal relationship between changes in LH secretion and GnRH messenger RNA (mRNA) expression at the time of the estradiol-induced LH surge. Ovariectomized Clun Forest ewes were treated with exogenous progesterone and estradiol (E) to mimic estrous cycle concentrations of these gonadal steroids and to induce the LH surge. Ewes were killed at five time points relative to the time of onset of the LH surge: pre-E, before E insertion (n = 6); presurge, after E insertion and 8–10 h before surge onset (n = 5); ascending limb, 2–6 h after surge onset (n = 5); midpeak, 9–12 h after surge onset (n = 5); and postsurge, 21–27 h after surge onset (n = 5). Control animals (n = 5/group), which received no E, were killed at identical time intervals alongside the E-treated ewes. Coronal sections containing the diagonal band of Broca through to the anterior hypothalamus were processed for cellular in situ hybridization using an 35S-labeled oligonucleotide probe complementary to ovine GnRH. No changes were found in the number of GnRH mRNA-expressing cells detected in the rostral preoptic area or the medial septum in either gonadal steroid-treated or control ewes. In contrast, cellular GnRH mRNA expression (as assessed by silver grain density) decreased significantly (P < 0.05) between presurge and ascending limb groups within both the rostral preoptic area (0.64 ± 0.06 vs. 0.43 ± 0.05 silver grain density/µm2) and medial septum cells (1.08 ± 0.09 vs. 0.77 ± 0.07). No significant changes were detected in control ewes. These results show that the estradiol-induced LH surge in the ewe is associated with a decrease in GnRH mRNA expression that occurs in advance of the onset of the GnRH surge. This suggests that neural mechanisms controlling GnRH biosynthesis may be distinct from those regulating GnRH secretion.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE PREOVULATORY LH surge is generated by increasing circulatory levels of estradiol (E), which stimulate a surge of GnRH release into the portal circulation (1, 2, 3, 4, 5) and maximize gonadotroph responsiveness to GnRH (6, 7, 8). The profile of GnRH secretion at the time of the surge has been well characterized in the ewe (3, 9, 10, 11). Thus, a gradual increase in GnRH pulse frequency and amplitude coupled with elevated GnRH interpulse secretion have been shown to begin approximately 3–4 h before the massive and sustained release of GnRH that generates the LH surge (3, 11). Although this pattern of secretion results in the release of substantial amounts of GnRH, the nature of any changes in GnRH biosynthesis in relation to the ovine surge is unknown.

As one index of GnRH biosynthesis, several investigators have examined the relationship between changes in GnRH messenger RNA (mRNA) expression and the LH surge in the rat. Although several studies conclude that temporally and regionally restricted increases in cellular GnRH mRNA content occur during the steroid-induced or proestrous LH surge (12, 13, 14, 15, 16), this point is still controversial (16, 17, 18). Where changes in GnRH mRNA have been reported, the precise temporal relationship between the onset of the changes in mRNA and the LH increment remains unclear. For example, some investigators (14, 15, 16) have demonstrated that GnRH mRNA content rises in the rostral preoptic area (rPOA) at least 2 h before LH levels are seen to rise, whereas others have shown that the number of detectable GnRH mRNA-expressing cells in the more caudal POA (14) or whole rPOA (13) increases with the onset of the LH surge. Hence, it remains unclear whether a common neural mechanism may operate to alter GnRH biosynthesis and GnRH secretion at the time of the preovulatory LH surge and what causal relationship exists between the increase in mRNA and secretion.

In the present study, we sought to determine whether changes in GnRH mRNA expression occur at the time of the GnRH/LH surge in the ewe and, if so, to establish the temporal relationship between GnRH biosynthesis and secretion. This animal model has particular advantages, in that under conditions of prior progesterone withdrawal, the GnRH surge is known to be initiated by rising E concentrations alone, and the profile of GnRH secretion leading up to and throughout the surge is well established. Hence, by using in situ hybridization to assess GnRH mRNA content, we have been able to examine changes in cellular GnRH transcript levels within anatomically distinct GnRH cell populations in relation to known fluctuations in GnRH secretion throughout the time of the preovulatory surge.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals and treatments
Fifty-one Clun Forest ewes were used in this study, which was conducted during the breeding season (October/November) at Babraham (Cambridge, UK; 52°, 12'N). During the study, ewes were kept indoors under natural lighting conditions and maintained on a diet of sheep concentrate and ad libitum hay and water.

Each ewe was ovariectomized and run through two artificial 14-day estrous cycles using a modification of the model of Goodman and colleagues (19). The artificial luteal phase was produced by the insertion of progesterone (1 x 9% progesterone CIDR-G intravaginal device, InterAg, Hamilton, New Zealand) and E (1 x 10 mm sc 17ß-estradiol SILASTIC brand implant, Dow Corning, Midland, MI) for 11 days. The follicular phase was initiated by the removal of progesterone, followed 16 h later by the insertion of 4 x 30-mm sc 17ß-estradiol SILASTIC implants (E).

During the first artificial follicular phase, LH concentrations were monitored in jugular blood from all ewes at hourly intervals from 8–40 h after E insertion. Data from this cycle were used to predict when a LH surge would occur in the second cycle. During the second artificial follicular phase, ewes were split into two groups and received E (surge treatment) or were subjected to sham implantation (control treatment). Ewes were killed at one of five time points relative to the time of predicted LH surge onset as follows: pre-E (n = 6), immediately before E insertion; presurge (n = 5), after E insertion but 8–10 h before LH surge onset; ascending limb (n = 8), 2–6 h after LH surge onset when LH levels are rising exponentially; midpeak (n = 7), 9–12 h after surge onset when LH levels are decreasing from the surge peak; and postsurge (n = 5), when LH levels have returned to baseline. Ewes receiving the control treatment were time matched relative to their predicted time of LH surge onset, and five animals were killed at each of the time points described above.

Hourly samples of jugular blood were taken from 6 h before E insertion until the time of death to assess LH secretion profiles. In addition, jugular blood samples were taken at half-hour intervals in both ascending and midpeak groups from 2 h before death to determine more precisely the pattern of LH release. Ewes were killed by barbiturate overdose (20 mg/kg BW, iv; Lethobarb, Duphar Vet, Southampton, UK), and the brain was rapidly removed. A block of tissue approximately 1.5 cm3 containing the preoptic area and hypothalamus was dissected from the brain within 3 min of death and rapidly frozen on dry ice. Tissue was stored at -70 C before sectioning. All animal procedures were conducted under a project license (PPL 80/1037) issued by the Home Office.

GnRH in situ hybridization
Frozen brain sections (15 µm thickness) were cut in the coronal plane from the level of the diagonal band of Broca to the anterior hypothalamus (AHA) on a cryostat (Bright, Huntingdon, UK). Brain sections were thaw-mounted onto Vectabond-coated slides (3 sections/slide) so that 10 sets of every 10th section taken from the diagonal band of Broca to the AHA were compiled for each animal. Sections were stored at -70 C until used.

A 39-mer oligonucleotide probe complementary to the GnRH-encoding region of the partially cloned ovine GnRH complementary DNA sequence (20) was used for GnRH mRNA in situ hybridization (probe sequence, 5'-TCT CTT TCC TCC AGG GCG CAG CCC ATA GGA CCA GTG CTG-3'). The six 3'-residues of the probe are complementary to those encoding the first two amino acids of GnRH and, although not yet identified in the sheep, are identical in human, mouse, and rat GnRH sequences. EMBL database searches showed no significant homology other than that of other mammalian GnRH coding sequences. The oligonucleotide was 3'-end labeled with [35S]deoxy-ATP (1000–1500 Ci/mmol; New England Nuclear-DuPont, Boston, MA) using terminal deoxynucleotidyl transferase (50 U; Pharmacia, Uppsala, Sweden) and purified by filtration on a Sephadex G-50 column, resulting in a specific activity of approximately 109 cpm/mg probe.

Because of the large number of slides involved (three or four slides per animal with five or six animals per group and five different time points), the in situ hybridization was undertaken as two separate experiments, 24 h apart. Slides from the five control group ewes underwent hybridization together as one set, whereas slides from the five E-treated groups of ewes were processed in an identical fashion the following day using the same labeled probe. Sections were quickly warmed to room temperature using a hair dryer, fixed with 4% paraformaldehyde in 0.1 M phosphate buffer for 20 min, and rinsed in 0.1 M PBS. Sections were dehydrated through increasing ethanol concentrations of 70%, 80%, 90%, 95%, and 100% and allowed to air-dry. For hybridization, the 35S-labeled probe was diluted in hybridization buffer [20 x saline sodium citrate (SSC), 50% deionized formamide, 10% dextran sulfate, 1 x Denhardt’s solution, 250 µg/ml sheared salmon testicular DNA, and 0.3% ß-mercaptoethanol] to give a final concentration of approximately 1.2 x 103 cpm/µl, and 250 µl were applied to each slide. After overnight hybridization at 37 C, sections were washed in 1 x SSC at room temperature, three times in 1 x SSC at 55 C (30 min each), and finally in 1 x SSC for 1 h at room temperature. After a brief rinse in water, sections were placed in a 300-mM ammonium acetate-70% ethanol solution for 30 sec, followed by absolute ethanol for 30 sec, then allowed to air-dry. The slides were assorted randomly, dipped in Ilford K-5 nuclear track emulsion, and exposed for 2–3 days in light-tight boxes. All slides were developed with Ilford Phenisol (diluted 1:5 in distilled water; 5 min at 20 C) and counterstained lightly with methylene blue. Competition experiments were undertaken in which the 35S-labeled probe was applied to the brain sections in the presence of a 50-fold excess of unlabeled probe.

Analysis
The number of cells expressing GnRH mRNA per section was determined for each animal by counting the total number of positively hybridized cells located within anatomically matched sections containing the rPOA and medial septum (MS). Cells were considered to be positively hybridized when silver grains were found clustered over a methylene blue-counterstained cell body, and the number of silver grains was greater than 5 times the number counted over preoptic cells in the excess cold oligo controls (21, 22). Hybridized cells were assigned to either the rPOA or MS depending on their anatomical location, as revealed by the methylene blue counterstaining. Specifically, all hybridized cells clustered around the organum vasculosum of the lamina terminalis (OVLT) and the rostral extension of the supraoptic recess of the third ventricle were classified as rPOA GnRH neurons (Fig. 3Go, B–D). Cells found rostral to the OVLT (Fig. 3AGo) were few in number and, therefore, were excluded from the analysis. Hybridized cells lying dorsal to this population, in the midline, and surrounded by the morphologically distinct magnocellular neurons of the MS, were termed MS GnRH neurons (Fig. 3Go, C and D). As insufficient material existed with which to accurately match all groups, the hybridized cells found more caudally in the ventrolateral aspect of the anterior hypothalamus were not analyzed. Cell counts from three or four sections for each GnRH population were used to provide individual animal means and then combined to provide treatment group means and SE values at each time point.



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  		Figure 3. Camera lucida diagram of GnRH mRNA hybridized cells through a series of rostro-caudal sections (A–D) in one individual ewe (each section is 300 µm apart). Small circles indicate cells analyzed as rPOA GnRH neurons, whereas large circles indicate cells analyzed as MS GnRH cells. Stars indicate cells lying outside the rPOA and MS that were not analyzed. 3v, Third ventricle; ac, anterior commissure; oc, optic chiasm.

 
An assessment of cellular GnRH mRNA content was made by analyzing silver grain density over individual cells using a Seescan Sonata II image analyzer (Seescan, Cambridge, UK) coupled to a Leica Orthoplan microscope as reported previously (21, 22). With this system the operator outlines the silver grain cluster over each cell, and silver grain density is determined in arbitrary units per µm2. The same anatomical criteria outlined above were used to distinguish rPOA from MS GnRH neurons, and all hybridized cells found in each area were analyzed. Individual values were combined to give an average for each animal, and these values then used to form group means and SE values at each time point. Because of the priority to compare mRNA content between the time point groups in defined populations, all sections were analyzed for the rPOA cells first, then the MS neurons were analyzed at a later time. A frequency distribution chart of relative silver grain density within the GnRH population was compiled by determining the percentage of neurons expressing silver grain densities in 0.1 arbitrary units per µm2 bins (i.e. 0–0.1, 0.1–0.2 ... ). These data were compiled for each ewe and combined to give time point group means and SE values.

Statistical analysis of cell numbers and silver grain density between time points within the control and E-treated experimental groups were assessed using ANOVA with Tukey’s post-hoc test. Significance was set at P < 0.05.

LH RIA
Plasma LH was measured in duplicate 100-µl aliquots using a previously described, double antibody RIA procedure with the anti-LH antiserum CSU 204 (G. D. Niswender, Fort Collins, CO) and the NIDDK S11 standard (23). Inter- and intraassay coefficients of variation were 13.24% and 8.67%, respectively, for six assays. The average detection limit of the assay was 0.195 ng/ml. The time of onset of the LH surge was defined as the first sample of a sustained rise in LH levels that was greater than the mean ± 2 SD of the five preceding samples.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
LH results
All 51 ewes exhibited a LH surge during the first artificial follicular phase. The mean time to surge onset was 16.4 ± 0.3 h (range, 12–23 h) after E implantation. Mean LH concentrations during the second artificial follicular phase are shown at each time point after control (Fig. 1Go, right panel) and surge treatments (Fig 2Go, left panel). Due to within-animal variation, not all animals in the ascending limb and midpeak groups were killed at the appropriate times. Ewes in these groups were reallocated, based on individual LH data, resulting in both the ascending limb and midpeak groups containing five animals each. The five remaining animals from these two groups were killed before the LH surge began and thus were excluded from analysis.



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  		Figure 1. Profiles of plasma LH leading up to death in the ewes that received E (surge group; left) and animals that received no E (control group; right) at each of the five time points chosen for GnRH mRNA analysis.

 


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  		Figure 2. Darkfield photomicrographs of silver grain clusters over cells in the rPOA (A) and MS (B) after hybridization with the 35S-labeled oligonucleotide probe complimentary to ovine GnRH. Note that A and B are at different magnifications. Scale bar = 100 µm. 3v, Third ventricle; ac, anterior commissure; oc, optic chiasm.

 
GnRH cell distribution
Individual hybridized cells were recognized by a cluster of reduced silver grains overlying the cell. The specificity of hybridization was demonstrated by the lack of similar silver grain deposition over individual cells in the presence of excess unlabeled oligonucleotide (data not shown). Hybridized cells were found located in a continuum throughout MS and POA/AHA region, with the largest numbers of hybridized cells located in the rPOA in close proximity to the supraoptic recess of the third ventricle, the optic chiasm, and the OVLT (Figs. 2AGo and 3Go, C and D). Smaller numbers of hybridized cells were also found in the midline above and rostral to the rPOA in the MS (Figs. 2BGo and 3Go, C and D).

Effects of E treatment on hybridized cell number
No significant differences (P > 0.05, by ANOVA) were evident in the number of hybridized cells detected in the rPOA or MS of control or E-treated group across the five time points (Table 1Go).


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  		Table 1. Mean number of cells ± SE hybridized with ovine GnRH mRNA oligonucleotide probe per section (rPOA, left column; MS, right column)

 
Effects of E treatment on cellular silver grain density
No differences in cellular silver grain density were detected in either the rPOA or MS across the five time points examined in the control ewes (Fig. 4Go). In contrast, a significant change in mean cellular silver grain density over the five time points was found in both the rPOA and MS of E-treated ewes (P < 0.05, by ANOVA). Specifically, a significant decrease in cellular silver grain density occurred between the presurge and ascending limb time points in both the rPOA and MS (rPOA cells, 0.64 ± 0.06 vs. 0.43 ± 0.05 silver grain density in arbitrary units/µm2; MS cells, 1.08 ± 0.09 vs. 0.77 ± 0.07; P < 0.05, by Tukey’s post-hoc test). This was preceded by a nonsignificant increase between pre-E and presurge time points. Silver grain density remained relatively constant over the ascending limb, midpeak, and postsurge time points in both rPOA and MS cells (Fig. 4Go).



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  		Figure 4. Mean cellular GnRH mRNA content (±SE) from individual ewes analyzed at each chosen time point after control (top panel) and experimental (bottom panel) treatments. Cells analyzed in the rPOA and MS are shown in the left and right panels, respectively. GnRH mRNA content decreased significantly between the presurge and ascending limb groups after surge treatment in both rPOA and MS cells. *, P < 0.05, by Tukey’s post-hoc ANOVA test.

 
The frequency distribution of silver grain density in rPOA cells of the E-treated ewes revealed an approximately normal distribution pattern of mRNA expression throughout the five time points, which reflected the changes in mean silver grain density (Fig. 5Go).



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  		Figure 5. Frequency distribution of GnRH mRNA expression within the rPOA population in the five experimental groups of ewes. Each graph depicts the mean percentage (±SE) of cells at different silver grain densities (0.1-µm2 silver grain density bins).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
To analyze GnRH mRNA expression in the sheep, an oligonucleotide probe specific for ovine GnRH was developed using the material published for the partially cloned ovine GnRH complementary DNA sequence (20) and knowledge of highly homologous sequences in mammalian GnRH DNA. Using this probe, we show here that the distribution of hybridized neurons is identical to that found by GnRH immunocytochemistry in the ewe (24, 25) and that the hybridization signal, achieved with a 2-day exposure time, was completely abolished in competition experiments with unlabeled probe. Together, these observations demonstrate the specificity of our method for detecting GnRH mRNA in the ovine brain.

This study has examined the temporal changes in cellular GnRH mRNA expression that are associated with the E-induced LH surge in the ewe. Our results show that significant changes in GnRH mRNA expression occur in ewes before the onset of the estradiol-induced LH surge, whereas time-matched control animals exhibit no changes. Specifically, cellular GnRH mRNA content was found to fall between the presurge and ascending limb time points of our study, which represents an approximately 8-h period of time over which circulating LH concentrations change from basal levels to the initial increment at the start of the LH surge. Recent work has shown that the latter portion of this period represents a time when a gradual increase in GnRH pulse frequency and amplitude coupled with elevated GnRH interpulse secretion occur before the onset of the GnRH surge (11). Hence, our findings indicate that in the ewe, GnRH mRNA levels fall over a period when GnRH secretion is increasing gradually before the onset of the GnRH surge.

It is also of interest to note that GnRH mRNA remained constant over the ascending limb, midpeak, and postsurge time points. Using an animal model similar to that used in this study, Moenter and colleagues (26) reported that Fos expression (a marker of neuronal activation) was increased in GnRH neurons during the GnRH surge. As the maximum number of Fos-positive neurons occurs after the initiation of the surge, these authors suggest that Fos induction may be related to the replenishment of GnRH in neurons requiring an increase in cellular GnRH mRNA. Our data do not support this view. Thus, it does not appear that the surge-related increase in Fos leads to an acute increase in cellular GnRH mRNA content.

Species comparisons
These observations in the sheep appear to be in contrast to those reported in the rat. Thus, where changes in GnRH mRNA expression are reported these constitute an increase in mRNA that is closely associated with the LH surge (12, 13, 14, 15, 16). In terms of the absence of an increase in cellular GnRH mRNA content in the ewe, we did note a nonsignificant increase in GnRH mRNA expression after estrogen administration in the presurge group before the significant fall in mRNA content. Hence, it is possible that a rise in GnRH mRNA expression also occurs in the ewe, but does so many hours before the preovulatory changes in GnRH secretion. Perhaps a larger sample size would have allowed us to make this determination. However, because of the necessity in this study to examine GnRH mRNA content in the ewe at time points separated by relatively long time intervals, it is possible that we may have missed a peak of GnRH mRNA expression. Future studies will focus on the time period following the administration of E, but before the GnRH surge begins.

In terms of the temporal relationship between the change in GnRH mRNA and the GnRH surge, there would appear to be a clear species difference. The reason for this is unclear. It is, however, worth noting that the sheep model used in these studies is more physiological than the E-induced LH surges produced in some of the rat studies. In the latter case the LH surge is reduced in amplitude and delayed compared with a surge produced by estrogen and progesterone. Other factors that may be relevant are the differences between rats and sheep in the length of the ovarian cycle, the lack of a true luteal phase in the rat, and the degree to which a circadian input is used to trigger the onset of the surge (27, 28). Hence, although we can only speculate about why the profile of GnRH mRNA changes leading up to the LH surge are different between rats and sheep, one possibility is that the circadian trigger plays a role in generating the mRNA increase as well as synchronizing it with the neural apparatus responsible for GnRH secretion at the time of the surge.

Relationship between GnRH secretion and biosynthesis
As the changes in mRNA biosynthesis do not occur concomitantly with the onset of GnRH surge secretion in the ewe, our results suggest that different mechanisms may underlie these phenomena. Although the GnRH surge clearly represents the stimulatory actions of E on the GnRH neurosecretory system, the reduction in GnRH mRNA that we have observed may be related more to the negative feedback effects of E on GnRH secretion that precede the GnRH surge in this species (10, 11). At the least, it would seem that the effects of estrogen on GnRH mRNA expression and secretion are uncoupled in the hours leading up to the surge as mRNA content falls while pulsatile and interpulse GnRH secretion increases. Although, the direction of change in GnRH mRNA and secretion is the same in the rat, the temporal differences between the onset of the two events reported by some investigators have similarly led them to suggest that the neural mechanisms underlying the secretory and mRNA changes in the rat may also be different (15).

As GnRH neurons in the ewe (29, 30) and other species (31, 32, 33) do not contain classical nuclear estrogen receptors, estrogen-receptive interneuron populations may play a role in transmitting the estrogen signal to GnRH neurons. Recent work has shown that estrogen receptor-containing neurons located in the arcuate nucleus project to the median eminence (34), whereas estrogen-receptive cells located in the POA as well as the mediobasal hypothalamus project to the vicinity of the GnRH cell bodies in the rPOA of the ewe (Herbison, A. E., and A. Caraty, unpublished observation). These observations suggest that estradiol may have the capacity to modulate GnRH biosynthesis and secretion independently through different neuronal populations that target the GnRH cell body or terminal, respectively. If so, these different neural pathways, which have only been partially defined in terms of their neurotransmitter content (35), may underlie the differential regulation of GnRH neurons by estrogen.

The mechanisms underlying the changes in GnRH mRNA identified in this study are not known. Although the influence of estrogen on GnRH mRNA content in the rat is known to involve increases in GnRH gene transcription (16, 36), there is increasing evidence that supports the posttranscriptional regulation of GnRH mRNA content in vivo (37). In the present study we have identified a decrease in GnRH mRNA content. As changes in the stability of GnRH mRNA can be relatively rapid (38, 39), it is possible that this might result from increased degradation of the message. Such a conclusion awaits further studies.

Conclusion
In summary, this study has evaluated the changes in GnRH mRNA expression associated with generation of the GnRH surge in the ewe. That a significant decrease in expression is reported before the GnRH surge suggests that changes in GnRH biosynthesis precede the massive and sustained release of GnRH in this species and that GnRH mRNA expression may be declining at a time when GnRH secretion increases. The significance of these changes in GnRH mRNA expression to the successful generation of the GnRH surge warrants further investigation.


   Acknowledgments
 
The authors thank Andrew Dady, Martin White, Sam Herbert, Mike Bacon, and Tony Jones for help with the large animal work; James Taylor and Sandra Dye for assistance with the RIA procedure; Ian King for his expertise with tissue histology; and Dr. John Bicknell for comments on a draft of this manuscript. We are also indebted to the NIDDK (Bethesda, MD) for supplying both the LH for iodination and the LH standard preparation.


   Footnotes
 
1 A preliminary report of this work was presented at the 22nd Annual Meeting of the Society for Neuroscience, Washington, D.C., 1996 (Abstract 533.5). This work was supported by the BBSRC. Back

2 Present address: Reproductive Sciences Program, University of Michigan, Ann Arbor, Michigan 48109. Medical Research Council-funded graduate student. Back

3 Present address: I.N.R.A., Physiologie de la Reproduction des Mammifères Domestiques, 37380 Nouzilly, France. Research Fellow of St. Catharine’s College (Cambridge, UK). Back

4 Lister Institute-Jenner Fellow. Back

Received July 3, 1997.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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