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Regular Article |
a MRC Human Reproductive Sciences Unit, Centre for Reproductive Biology, Edinburgh EH3 9ET, United Kingdom
ABSTRACT
This study investigated whether prolactin (PRL) plays a priming role in the testis during the nonmating season and thereby facilitates gonadal reactivation. Sexually inactive Soay rams under long days were treated as follows: 1) group C (control) received vehicle, 2) group B received bromocriptine to suppress PRL secretion, 3) group B + PRL received bromocriptine + ovine PRL to reinstate physiological levels of PRL (n = 5/group). Treatments were for 10 wk. The photoperiod was then switched to short days to reactivate the reproductive axis. Testis diameter and sex skin coloration were recorded, and routine blood samples were collected to measure concentrations of FSH, inhibin A, and testosterone (T). At the end of the treatments, blood samples were collected every 10 min for 10 h to monitor LH pulses and the T-response to exogenous LH, and a testis biopsy was collected to assess spermatogenic activity (bromodeoxyuridine [BrDU] method) and expression of PRL receptor (reverse transcription-polymerase chain reaction and immunocytochemistry). There were no significant differences between groups in spermatogenesis (BrDU index) or steroidogenesis (T-response), and no difference in the time taken to achieve full testicular redevelopment under short days. Testis diameter and inhibin A were marginally increased in group B + PRL. Overall, this thorough experiment provides minimal support for the priming hypothesis.
anterior pituitary, FSH, GnRH, inhibin, LH, prolactin, seasonal reproduction, spermatogenesis, testes, testosterone
INTRODUCTION
Experimental studies in sheep have provided clear evidence that prolactin (PRL) acts directly in the testis, in concert with the classical gonadotropins LH and FSH, to regulate the seasonal testicular cycle in the adult ram. For example, treatment of rams with bromocriptine during summer, to chronically suppress PRL secretion, decreases testicular volume, sperm production, and testosterone secretion, and may delay testicular recrudescence in autumn [1–3]. In rams in which gonadotropin secretion is blocked permanently by surgical disconnection of the pituitary gland (HPD rams), alterations in photoperiod induced cycles in PRL secretion and parallel changes in testicular size with a 4- to 8-wk time-lag, consistent with a long-term stimulatory effect of PRL in the testis [4]. Furthermore, recent studies have demonstrated expression of PRL receptor (PRL-R) in the testis of both control and HPD rams, localized to Leydig cells in the interstitial tissue, and to germ cells undergoing spermatogenesis in the seminiferous tubules. Incubation of testis explants with PRL induced rapid phosphorylation of second messenger proteins (Jak2 and Stat1), providing evidence of a functional PRL-R in ovine testicular tissue [5].
Overall, these results support the priming hypothesis originally suggested by Sanford and Dickson [1]. This proposes that PRL acts in the testis under long days (LD) in spring and summer (when PRL secretion is markedly increased), and this facilitates gonadal reactivation induced by short days (SD) in autumn. The progonadal effect of PRL is presumed to be similar to that demonstrated in seasonal rodents [6, 7]. In sheep, however, there is an inverse relationship between PRL and gonadotropin secretion induced by photoperiod [8], and this creates an apparent paradox: PRL appears to promote testicular activity in rams at the nadir of the reproductive cycle.
The purpose of the current study was to test experimentally the priming hypothesis. The strategy was to house animals under LD to induce regression of the reproductive axis and to use a chronic 10-wk treatment with bromocriptine, with and without PRL replacement therapy, to establish the effects of PRL per se on the reproductive axis. The predictions were that PRL would affect one or more of the following: 1) expression of PRL-R in the testis, 2) steroidogenic and spermatogenic activity in the testis, 3) rate of reactivation of the testis induced by SD, and 4) short- and long-term patterns of gonadotropin secretion. The experimental protocol was designed to investigate each of the four predictions. Dual bromocriptine/PRL treatments have been used previously to demonstrate the role of PRL in the control of the seasonal pelage cycle in the Siberian hamster [9] and cashmere goat [10], and to investigate the involvement of PRL in photoperiod-induced body weight cycles in sheep [11].
MATERIALS AND METHODS
Animals
The study was conducted under a Project Licence issued by the Home Office and adhered to strict ethical guidelines for animal experimentation. Adult rams of the Soay breed of feral sheep that show pronounced photoperiod-induced cycles in testicular activity, coat growth, and other seasonal characteristics were used in the study [12]. The animals were adult (2 yr old) with a mean body weight of 34.3 kg (range 31–39 kg) at the start of experiment. They were housed permanently in light-controlled rooms and routinely exposed to alternating 16-wk periods of long days (16L:8D, LD) and short days (8L:16D, SD) to entrain the testicular cycle. Light intensity was approximately 160 lux at the animals' eye level. The time of lights-on was constant (0800 h), and adjustments in photoperiod were achieved by abruptly changing the time of lights-out by 8 h. The animals received a maintenance diet of commercial grass nuts (Vitagrass; Vitagrass Farms Ltd, Cumbria, UK), with hay and water ad libitum. Ambient temperature in the animal rooms was maintained between 10 and 20°C.
Photoperiod Manipulations and Hormone Treatments
The experimental treatments were initiated at 8 wk into LD when the rams were sexually inactive with high circulating concentrations of PRL. Fifteen animals were assigned to three equal groups balanced for body weight (n = 5/group). These were treated as follows: 1) group C (control) received the vehicle treatment as for the bromocriptine group for 10 wk, 2) group B received bromocriptine for 10 wk (daily dose, 2 mg/ram), and 3) group B + PRL received bromocriptine (as above) and ovine PRL (oPRL) for 10 wk (daily dose, 10 mg/ram). All groups were maintained on LD during the treatments and transferred immediately to SD after the treatments. The total experimental period was 32 wk (8 wk pretreatment, 10 wk treatment, 14 wk post-treatment); this allowed enough time for full regression and reactivation of the testis in all groups.
The bromocriptine (2-bromo-
-ergocriptine; Sigma-Aldrich, Poole, Dorset, UK) was prepared daily, dissolved in ethanol, and then diluted with 0.9% saline (50:50 v:v; vehicle) to produce a 1-mg/ml solution. Each ram in group B and B + PRL received 1.0 ml solution subcutaneously on the side of the neck twice daily (1 mg bromocriptine at 0800 and 2000 h). The site of injection was varied to prevent local inflammation. The daily dose of 2 mg/ram was selected because it had been shown to fully suppress endogenous PRL secretion [3]. The control rams received the vehicle injections at the same time as the experimental groups.
The oPRL (3.5 g, LER-1790) was purified from sheep pituitary glands (Professor L.E. Reichert, Department of Biochemistry and Molecular Biology, Albany Medical College, NY). The potency, determined by the pigeon crop sac assay, was 26.4 U/mg (95% confidence limits = 17.9–38.9). Contamination with FSH was <0.020 NIH-FSH-S1 U/mg (hCG-augmented ovarian weight-gain method), with LH was <0.004 NIH-LH-S1 U/mg (ovarian ascorbic acid-depletion assay), with thyroid-stimulating hormone (TSH) was 0.004 NIH-TSH-S1 U/mg, and with growth hormone was <0.010 IU/mg (hypox rat weight-gain assay; L.E. Reichert, personal communication). Aliquots of freeze-dried PRL (50 mg) were dissolved daily in 12 ml 0.9% saline + 150 µl 0.2 M NaOH. Once in solution, the pH was adjusted to near neutrality by the careful addition of 0.2 M HCl. Each ram in group B + PRL received 1.0 ml solution s.c. on the side of the neck two times daily (5 mg oPRL at 0800 and 2000 h). The sites of injection were again varied to prevent local inflammation. The daily dose of 10 mg/ram was selected to maintain circulating concentrations of PRL in the physiological range for rams living under LD [13]; this was confirmed in a preliminary trial. The animals tolerated the multiple injections with no clear side effects other than hardening of skin.
Routine Measurements
To record the long-term endocrine changes during the experiment, blood samples were collected from each animal by venipuncture of the jugular vein. This was carried out twice weekly during the early light phase (approximately 2 h after the morning routine injections). The blood samples were heparinized, and the plasma separated within 30 min and stored at -20°C. Every 2 wk the diameter of the testis was measured within the scrotum using calipers, and the intensity of the sexual skin coloration in the inguinal region was visually scored, as an immediate index of gonadal activity [14]. The pattern of growth (short, <0.6 cm; medium, 0.7–1.4 cm; long, >1.5 cm) and molting of the pelage on the scrotum (% molt/ease of plucking) was also recorded as a biological response to PRL [13].
Collection of Serial Blood Samples and a Testis Biopsy
On one occasion at the end of the treatment phase (Week 18), blood samples were collected from the jugular vein every 10 min for 10 h to monitor pulsatile LH secretion. The sampling started in the early light phase (0830 h) close to the time of the routine injections of bromocriptine and PRL. After 8 h each animal received a bolus injection of oLH (4.0 µg/ram i.v. NIADDK-oLH-26, Bethesda, MD), to record the effect on the secretion of testosterone (index of testicular steroidogenesis). The dose of LH was selected to produce a peak in circulating concentrations of LH similar to, or marginally greater than, the peak concentration associated with an endogenous LH pulse during the reactivation phase of the reproductive cycle [15]. To facilitate the repeated sampling, a poythene cannula was inserted into the jugular vein on the day before study and was kept patent with heparinized saline (10 000 IU heparin/1.0 L 0.9% saline). The blood samples were collected into heparinized tubes on water ice, and the plasma was separated within 30 min and stored at -20°C.
A testicular biopsy was also collected from each animal at the end of the treatment phase (Week 18, 2 days after the collection of serial blood samples). The animals were pretreated with 5-bromo-2-deoxyuridine (BrDU; 5 mg/kg i.v.), given 2 h before the biopsy, to characterize the dividing cells within the testis [16, 17]. The BrDU was dissolved in warm 0.9% saline shortly before use and given at a concentration of 20 mg/ml. The testis biopsy was removed under local anaesthetic. Small fragments of testicular tissue (approximately 5 x 2 x 2 mm) were fixed in Bouin solution for 6 h and stored in 70% ethanol before processing for immunocytochemistry. Tissue samples were also snap frozen in liquid nitrogen and stored at -70°C for the extraction of mRNA.
Radioimmunoassays
The concentrations of PRL, FSH, and LH in the blood plasma samples were measured using routine RIAs, validated for sheep plasma for PRL [18], FSH, and LH [19]. The PRL assay had a lower limit of detection (10% decrease in binding relative to Bo) of 0.5 µg/L NIH-PRL-S13 plasma, and an intra- and interassay coefficient of variation (CV) of 7.0 and 9.0%, respectively, based on low, medium, and high quality control samples measured in 14 assays. The corresponding values for the FSH assay were 0.2 µg NIDDK-FSH-RP2/L, 10.0% and 13.0%, and for the LH assay were 0.2 µg NIH-LH-S18/L, 5.5% and 8.0%, respectively. All samples from a single animal for a similar phase of the experiment were measured in the same assay.
The concentrations of testosterone in selected weekly samples and in serial samples collected after the injection of oLH were measured by RIA, with solvent extraction, and the use of iodinated tracer [20]. The sensitivity of the assay was 0.3 nM/L, and the mean intra- and interassay CV were <12%.
The concentrations of inhibin A subunit in selected weekly samples were measured using a specific two-site enzyme immunoassay [21]. As a standard, this assay used a highly purified preparation of human inhibin A, a monoclonal capture antibody (INPRO) against a sequence of the beta A subunit, and the same detection antibody (R1) as used in dimeric inhibin assays. Alkaline phosphatase activity was measured using pNPP substrate. The cross reaction with recombinant forms of inhibin B and follistatin was <0.02%, as demonstrated by immunoblotting studies. The detection limit was 3 µg/L.
RNA Extraction and Reverse Transcription-Polymerase Chain Reaction
To confirm the expression of PRL receptor in the testis biopsies, total RNA was extracted from the testicular tissue using the guanidinium thiocyanate method as previously described by Chomczybski and Sacchi [22]. RNA (1 µg) was treated with DNase 1 (amplification grade; Life Technologies, Paisley, UK), and cDNA synthesized using Expand reverse transcriptase (Roche, Lewes, UK). Oligonucleotides (sense 5'-CTG GTT GGT TCA TTA TCC AGT ACG-3' and antisense 5'-GCA GGT CAC CAT GCT ATA GCC CTT-3') were designed from the oPRL-R sequence [23], to generate a 312-base pair (bp) product (nucleotides 482–498). Polymerase chain reaction (PCR) was performed at an annealing temperature of 57°C for 35 cycles in 75 mM Tris/HCl (pH 9.0), 20 mM (NH4)2SO4, 1.5 mM MgCl2, 200 µM each deoxynucleotide-5'-phosphate, and 0.01% Tween-20 with 25 pmol of each primer and 1.25 U AGS Gold DNA polymerase (Hybaid, Ashfield, UK) in a final volume of 50 µl using 5 µl sheep testis cDNA as a template for the reaction. A separate PCR reaction was also run under the same conditions using glutaraldehyde 3-phosphate dehydrogenase (GAPDH) primers (450-bp product; sense 5'-ACC ACA GTC CAT GCC ATC AC-3' and antisense 5'-TCC ACC ACC CTG TTG CTG TA-3') as a positive control to check uniformity of the cDNA synthesis. After amplification, PCR products (8 µl) were electrophoresed through a 2% agarose gel containing 0.5% ethidium bromide and the products visualized and photographed under UV light. Samples from all animals in the three treatment groups were run in parallel for comparison.
Histology and Immunocytochemistry
To localize the PRL-R protein within the testicular tissue, the Bouin-fixed testis biopsies were progressively dehydrated through ethanol and embedded in paraffin wax. Sections (5 µm) were cut and mounted on slides coated with 2% 3-aminopropyltriethoxy-silane (TESPA) in acetone. Slides were then dried overnight at 50°C before dewaxing in Histoclear (National Diagnostics, Hull, UK). Tissues were rehydrated in graded ethanol and washed in water followed by PBS (0.05 M Tris-HCl, pH 7.4, 0.85% NaCl). Sections were treated with 10% hydrogen peroxide in methanol for 30 min and then blocked for 30 min with normal swine serum (NSS) diluted 1:5 in Tris-buffered saline (TBS) + 5% BSA. The primary antibody for the PRL-R (kindly donated by Dr. P.M. Ingleton, School of Medicine, University of Sheffield) was raised against a 16-amino acid synthetic peptide corresponding to residues 53–68 of the external domain of the rat PRL-R [23]. This sequence shows close homology with the corresponding ovine sequence of the PRL-R [24]. The receptor antibody had been validated for use in immunocytochemistry in sheep tissues using several control procedures including preadsorption of the primary antibody using the specific peptide sequence as used in the initial immunization [25] and had been used previously to localize PRL-R in the Soay sheep testis [5]. The polyclonal antibody was diluted in NSS-TBS plus 5% BSA (see above) and incubated on sections overnight at 4°C under plastic coverslips. Negative control sections were incubated with nonimmune rabbit serum. After removal of the coverslips, sections were washed twice in TBS (5 min each) and incubated for 30 min in biotinylated swine anti-rabbit immunoglobulin (Dako, High Wycombe, Bucks, UK) diluted 1:500 in NSS-TBS. The sections were then washed again twice in TBS (5 min each) and incubated with peroxidase anti peroxidase-conjugated to avidin-biotin complex (Dako) for 30 min at room temperature. Color reaction was developed by incubation in a mixture of 0.05% 3-diaminobenzidine (DAB; Sigma) in 10 ml 0.05 M Tris-HCl buffer (pH 7.4) and 0.033% hydrogen peroxide. Hematoxylin was used as a counterstain to help visualize the structure of the testicular tissue. Sections from animals in the three treatment groups were run in parallel to provide a semiquantitative comparison.
The BrDU-labeled proliferating cells in the testis samples were visualized using a mouse monoclonal antibody to BrDU (Boehringer Mannheim). Sections were dewaxed in Histoclear and rehydrated in decreasing concentrations of methylated spirits. Antigen retrieval was accomplished by microwaving sections for 4 x 5 min in 0.01 M citrate buffer. Sections were treated for 30 min with normal rabbit serum block (NRS; Dako), then incubated in mouse monoclonal anti-BrDU (3 µg/ml in TBS) at 4°C overnight. Control sections were treated with mouse IgG (Vector Laboratories, Peterbough, UK) (3 µg/ml in TBS) in place of primary antibody. Secondary antibody, rabbit anti-mouse (26 µg/ml in blocking serum), was applied for 30 min at room temperature. BrDU binding was visualized using mouse alkaline phosphatase/antialkaline phosphatase (APAAP; Sigma) (1 µg/ml in blocking serum), incubation for 30 min, and the nitroblue tetrazolium (NBT) detection system (Boehringer Mannheim). Sections were briefly counterstained with hematoxylin and mounted for analysis. To measure the level of cell proliferation within the seminiferous tubules the number of BrDU-positive cells in 50 tubules were counted. Tubules presented in cross section on the slide (round tubule profile) were selected at random in each animal.
Statistical Analysis
The significance of the difference between groups in plasma concentrations of PRL and FSH, and testis diameter during the treatment phase (Experimental Weeks 8–18) and during the post-treatment phase (Experimental Weeks 18–32) was assessed using two-way ANOVA with repeated measures. When a time x treatment interaction was significant (P < 0.001), the weeks when the values differed between groups were further analyzed using ANOVA. To summarize the changes in plasma hormone concentrations (FSH, inhibin A, and testosterone) in response to the change from LD to SD, the experiment was divided into four phases: phase 1, end of treatments under LD (Weeks 14–18), phase 2, early post-treatment under SD (Weeks 18–22), phase 3, mid post-treatment under SD (Weeks 22–26), and phase 4, late post-treatment under SD (Weeks 26–30; Fig. 3). To test for significant differences between groups, the values during each phase were compared using ANOVA with repeated measures. The mean hormone concentration was calculated for each phase for each animal, and this was used to calculate the mean ± SEM for each group (Fig. 5, significant difference indicated by letters). To test for significant differences with time, the values during each phase were compared within group C using ANOVA (Fig. 5, significant difference indicated by asterisk). The times of maximum testis diameter and plasma concentrations of FSH and testosterone following the switch to SD were assessed based on a three-point moving average for each parameter. This was expressed as weeks (mean ± SEM) from the light-change (Experimental Week 18). For the analysis of pulsatile LH secretion based on 10-min blood samples, a significant LH pulse was defined as two consecutive high values, at least one of which exceeded the mean of the preceding two values by twice the intra-assay CV [26]. This method has been shown to detect physiological LH pulses consistently as evidenced by correlated increases in plasma testosterone concentrations [27]. The amplitude of the LH pulse was calculated as the peak concentration, minus the mean of the two preceding baseline values. The mean pulse-amplitude was determined for each individual profile and this was used to calculate the group mean ± SEM. The testosterone-response to the injection of oLH was calculated as the mean concentration of testosterone from 0 to 120 min after the treatment, minus the concentration of testosterone in the pretreatment sample. The significance of the difference between groups was assessed by ANOVA or Mann-Whitney rank test for LH pulse frequency.
RESULTS
Expression of PRL-R in the Testis
Reverse transcription (RT)-PCR analysis of RNA extracted from the testis biopsies collected at the end of the treatment phase (Week 18) confirmed expression of PRL-R in the sheep testis (Fig. 1). The procedure amplified a 312-bp product that would be common to both the short and long forms of the PRL-R. A product was seen in all animals, and there was no difference in the apparent level of expression in the PRL-R mRNA between the rams in groups C, B, and B + PRL.
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The localization of PRL-R protein in the testis biopsies collected at the end of the treatment phase (Week 18) was investigated by immunocytochemistry, and a typical result for a single animal in each treatment is presented (Fig. 2). Negative control slides showed no staining (Fig. 2f, inset). Fully processed slides showed immunostaining clearly evident in most Leydig cells and in some larger blood vessels within the interstitial compartment. The cytoplasm of germ cells within the seminiferous tubules was also heavily stained indicating the presence of PRL-R protein in the germ cells from the primary spermatocyte until the elongating spermatid stage of spermatogenesis. There was no clearly defined difference between the three treatment groups in the level of immunostaining for the PRL receptor in the testis (Fig. 2).
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Spermatogenesis in the Testis
Bromodeoxyuridine was administered 2 h before the collection of the testis biopsies (Week 18) to identify proliferation of cells in the testis (Fig. 2). Primary spermatocytes in the germinal epithelium on the periphery of the seminiferous tubules were distinctly labeled as replicating cells. Cell counts revealed no significant difference between the treatment groups in this index of spermatogenesis (labeled nuclei/tubule cross section: 24.5 ± 4.0, 25.1 ± 5.0, and 19.4 ± 4.6 nuclei, mean ± SEM for groups C, B, and B + PRL, respectively, NS).
Long-Term Changes Throughout the Experiment
Prolactin Following the pretreatment period (Weeks 0–8), all animals had increased blood concentrations of PRL due to exposure to LD. During the treatments (Weeks 8–18), PRL concentrations were suppressed in group B and increased in group B + PRL (Fig. 3). The statistical analysis of the PRL profiles indicated a highly significant (P < 0.001) time x treatment interaction in the PRL concentrations. PRL values were significantly (P < 0.001) reduced in group B compared with groups B + PRL and C from Week 9 to 18, and significantly (P < 0.01) increased in group B + PRL compared with group C, on most occasions from Week 11 to 18. The marked difference (>40-fold) in PRL concentrations between group B and group B + PRL throughout the 10-wk treatment provided proof of the successful chronic hormone replacement regimen (Fig. 3). After the end of the treatments, and following transfer to SD (Week 18), the blood PRL concentrations decreased in all groups. Except for a transitory rebound increase in PRL levels in group B, PRL concentrations remained low in all three groups throughout exposure to SD (Fig. 3).
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FSH and testis size Following the initial period under LD, blood FSH concentrations were low and all animals were sexually inactive with regressing testes (Fig. 3). During the treatments, FSH levels remained markedly suppressed and testis diameter declined to a minimum by Week 16. The statistical analysis revealed that there was no significant difference in the FSH profiles between the three treatment groups between Weeks 8 and 18. The data for testis diameter revealed a significant (P < 0.001) effect of time and a significant (P < 0.05) time x treatment interaction. Testis diameter was significantly (P < 0.05) increased in group B + PRL compared with group B at Weeks 14 and 16, but not different from the controls.
After the end of the treatments and transfer to SD (Week 18), blood FSH concentrations increased markedly in all groups and there was a parallel reactivation of the testes and the reappearance of the androgen-dependent sex skin coloration. Peak FSH concentrations occurred at Weeks 26–28, and the maximum in testis diameter occurred at Week 30. The statistical analysis revealed a significant (P < 0.001) effect of time, but no time x treatment interaction in the changes in FSH concentrations and testis diameter. Peak concentrations of FSH, however, were notably higher in group B compared with group B + PRL, but with considerable variation between animals. There were no significant difference between the groups in the time to maximum FSH concentration (time after transfer to SD: 8.3 ± 0.5, 8.1 ± 0.6, and 9.3 ± 0.7 wk, groups C, B, and B + PRL, respectively, NS), the time to maximum testis diameter (11.2 ± 0.7, 11.6 ± 0.6 = 5; and 11.0 ± 0.4 wk, groups C, B, and B + PRL, respectively, NS), time to maximum sex skin coloration (9.7 ± 0.3, 10.0 ± 0, and 10.2 ± 0.2 wk, groups C, B, and B + PRL, respectively, NS), or in the mature size of the testis (testis diameter: 55.0 ± 0.8, 54.2 ± 0.7, and 54.8 ± 0.6 mm, groups C, B, and B + PRL, respectively, NS). More details of the hormonal response to SD are given below.
Pulsatile LH secretion The results for the serial blood collection at the end of the treatment (10-min intervals for 10 h at Week 18) are summarized in Figure 4. LH pulses were identified at low frequency (1–4 pulses/8 h), indicative of a suppressed reproductive state. Although PRL concentrations were markedly different between the treatment groups (confirming the results based on the biweekly blood samples), there were no differences in LH mean concentration, LH pulse frequency, and LH pulse amplitude (1.02 ± 0.29, 1.21 ± 0.23, and 1.39 ± 0.52 µg/L LH for groups C, B, and B + PRL, respectively, NS). The testosterone-response to the standard injection of 4 µg oLH was also similar for the three groups. This injection produced a peak in blood concentrations of LH of 17.1 ± 1.9 µg/L (mean ± SEM) after 10 min; this was 2–10 times the concentration associated with the endogenous LH pulses in the sexually inactive animals. The LH injection provoked an increase in the blood concentrations of testosterone that was evident at 20 min, and that reached a maximum at 40–80 min, with no difference in the profiles between groups (data not shown). Depending on treatment, the experimental animals received injections of bromocriptine (group B), or bromocriptine and oPRL (group B + PRL) during the 10-h serial collection of blood samples. Blood PRL concentrations were permanently suppressed in group B. In group B + PRL, concentrations were similar to control animals before the injection of PRL and reached a maximum (up to twofold greater than controls) 3–4 h after injection of PRL. This confirmed that the PRL replacement therapy was effective at permanently maintaining PRL concentrations within or above physiological LD values in this experimental group.
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Pelage cycle The pretreatment under LD induced molting of the pelage on the scrotum from Week 6 onward. In the rams treated with bromocriptine (group B), the molt was arrested and some coarse hair was retained. The texture of the skin was dry. In the animals supplemented with PRL (group B + PRL), the molt was rapidly completed and there was regrowth of fine, supple hairs as in the vehicle-treated controls. During exposure to SD, a longer coarse hair characteristic of the winter pelage was developed and the coat was again similar for the treatment groups by the end of the study.
Reproductive Response to SD
The changes in the concentrations of FSH, inhibin A, and testosterone at the end of the treatments and following transfer to SD are summarized in Figure 5 (phases 1–4, Fig. 3). The statistical analysis of the data for the control group (group C) revealed a significant (P < 0.01) increase in all three parameters during reactivation of the reproductive axis induced by SD. FSH concentrations were at a maximum at Week 24 (phase 3) and decreased again by Week 28 (phase 4), as the testes became fully active. This decrease was associated with a significant (P < 0.01) increase in testosterone, and occurred when inhibin A values were already at a maximum. The overall pattern was similar in groups B and B + PRL. However, there were some significant (P < 0.05) differences between the treatment groups in inhibin A and testosterone during the reproductive transition. Inhibin A concentrations were reduced in group B, compared with group B + PRL and group C at Weeks 20 and 24. Testosterone concentrations were reduced in group B + PRL compared with group C at Week 20, and reduced in group B compared with group C at Week 24.
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DISCUSSION
This study clearly demonstrates that the chronic manipulation of circulating PRL concentrations in the ram had minimal effect on the photoperiod-induced testicular cycle in the ram. The suppression or replacement of PRL over a period of 10 wk under LD produced no demonstrable effect on testicular steroidogenesis (basal and LH-induced testosterone release) or spermatogenesis (number of BrDU-labeled germ cells), and only a marginal effect on testicular size. Furthermore, the treatments had no effect on the time taken for the testes to become fully reactivated under SD. Therefore the priming hypothesis is not supported by these results.
The failure to demonstrate a clear effect of PRL was not due to the absence of PRL-R expression in the testis. Expression of PRL-R was demonstrated by RT-PCR in testis biopsies recovered from all animals, and PRL-R protein was localized to the Leydig cells and germ cells as assessed by immunocytochemistry. This pattern was similar to that described previously in the ovine testis [5] and appeared to be unaffected by the ambient concentration of PRL. The chronic manipulation of circulating concentrations of PRL was also successful. The treatment with bromocriptine caused a consistent suppression of endogenous PRL secretion during the treatment only, as predicted based on previous studies [3]. The exogenous PRL therapy in conjunction with bromocriptine, produced circulating concentrations within, or temporally above, the physiological range for untreated rams under LD [28]. Moreover, this replacement therapy was effective at inducing molting and growth of the summer pelage that is known to be a biological response to PRL [10, 13, 29]. This confirmed the biological potency of the exogenous PRL preparation. The animals receiving PRL (group B + PRL) had circulating concentrations of PRL measured by RIA some 40-fold higher than in the corresponding negative control group (group B). This difference was maintained for 10 wk allowing adequate time for PRL to affect testicular function, even if the responses had a long latency [4].
The lack of a clear gonadal response to PRL is in contrast to the marked and rapid effects induced by altering gonadotropin secretion. This was obvious following the switch to SD where the increase in circulating FSH concentrations was associated with rapid growth of the testis over a period of 12 wk. This change had commenced at the end of LD, which indicates that the animals were becoming refractory to the inhibitory effect of the LD after 18 wk. The reactivation of the reproductive axis is known to result from an increase in pulsatile GnRH secretion from the hypothalamus. This stimulates the synthesis and release of FSH and LH, and thus activates the testis [12, 30–32]. This effect can be induced in HPD rams by the pulsatile administration of GnRH and the effect on the testis is evident within 3 wk [33]. In the current study, LH pulses of low frequency were detected in the periperal blood at the end of the treatment phase under LD reflecting a low level of pulsatile GnRH secretion. There were no differences in the LH profiles between the experimental groups, thus indicating that the administration of bromocriptine ± PRL had no apparent central effect on GnRH release (based LH pulse frequency), and no effect at the level of the pituitary (based on LH pulse amplitude). This was a predicted possibility because dopaminergic neural pathways play an inhibitory role in the regulation of GnRH/gonadotropin secretion [34, 35], and PRL-R are expressed in gonadotropes [25]. Besides the effects of increases in gonadotropin secretion, a decrease in gonadotropin secretion at the nadir of the reproductive cycle causes a further decline in testicular activity. This has been demonstrated in HPD rams where permanent blockade of gonadotropin release induces a decrease in testis size and spermatogenic activty, to values well below those observed in sexually inactive control animals [4]. This indicates that the low circulating concentrations of FSH and LH present under LD still play a role in maintaining basal testicular activity during the sexually inactive phase of the reproductive cycle.
This evidence supports the view that changes in gonadotropin secretion predominantly regulate seasonal reproductive cycle, and the actions of PRL in the testis are tightly modulated by the prevailing gonadotropin environment. This fits with the current observation that manipulating PRL produces demonstrable effects only during the reproductive transitions. In this study, PRL treatment appeared to advance inhibin A secretion marginally when the testes were beginning to redevelop, and this was possibly associated with lower concentrations of FSH when compared with the appropriate control. PRL may thus act in concert with FSH, to promote the secretion of inhibins from the Sertoli cell, and thus influence the negative feedback regulation of FSH secretion at the level of the pituitary gland. Inhibin A, but not inhibin B has been measured in peripheral blood in the ram [36], and inhibin A appears to act along with testosterone to achieve homeostasis [37]. This is consistent with the sequence of events in the current study, where the decline in FSH concentrations after reactivation of the testes occurred coincident with the increase in testosterone secretion, and at a time when the inhibin A concentrations were already at a maximum. Because the PRL-R is expressed in germ cells and not in Sertoli cells, the positive effect of PRL on inhibin production may be mediated through a germ cell/Sertoli cell interaction. There was also evidence for minor differences in testosterone secretion in the rams during the transition that might indicate subtle effects of PRL in Leydig cells where the expression of PRL-R was demonstrated.
One final point to consider is whether the effects of PRL in the testis might be more evident in domesticated breeds of sheep with a less extreme seasonality of reproduction, compared with the feral Soay breed. In Merino, Dorset, Suffolk, and Ile de France breeds for example, rams show a less pronounced decline in testis size during the nonmating season [15, 38], and it is in these breeds that suppression of PRL secretion in summer using bromocriptine has been shown to decrease testis size [1–3]. This may indicate that in breeds where photoperiodic regulation of reproduction through control of gonadotropins is less dominant, other factors such as nutrition have a greater influence on the function of the testis [39]. In this situation, PRL may have a greater overall influence on the seasonal regulation of the testis.
In conclusion, this study demonstrates for the first time that PRL-R mRNA and protein are expressed in the ovine testis when PRL is chronically suppressed, or when PRL circulates in high physiological concentrations, as occurs in summer under LD. The PRL-R are expressed in germ cells and Leydig cells as described previously, but also in blood vessels within the testis. Chronic manipulation of PRL, however, failed to affect spermatogenesis or testicular steroidogenesis in the sexually inactive testis and had a minimal effect on the programming of the testicular cycle. The results support the view that in this highly seasonal model, gonadotropin secretion dominantly regulates seasonal reproductive cycle, and the priming effects of PRL in the testis are not apparent.
ACKNOWLEDGMENTS
We are grateful to Norah Anderson, Marjorie Thomson, and staff at the Marshall Building who collected the blood samples and provided the long-term care of the animals; to Ian Swanston, Vivian Grant, and Irene Cooper for the expert technical assistance with the hormone assays; to Chris McKinnell for expert help with the BrDU immunocytochemistry; and to Tom McFetters and Ted Pinner for the art work. The purified preparations of FSH and PRL were provided generously by the National Hormone and Pituitary Program, the National Institute of Diabetes and Digestive and Kidney Diseases, the National Institute of Child Health and Human Development, and the U.S. Department of Agriculture for use in the RIAs.
FOOTNOTES
1 Correspondence: G.A. Lincoln, MRC Human Reproductive Sciences
Unit, Centre for Reproductive Biology, 37 Chalmers Street, Edinburgh EH3
9ET, UK. FAX: 44 0 131 228 5571; g.lincoln{at}hrsu.mrc.ac.uk 
Accepted: May 4, 2001.
Received: March 8, 2001.
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