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Prince Henry’s Institute of Medical Research Block E, Level 4, Monash Medical Centre, 246 Clayton Road, Clayton, Victoria 3168, Australia
1 Department of Anatomy and Cell Biology, Monash University, Clayton, Victoria 3168, Australia
(Requests for offprints should be addressed to Sarah Meachem; Email: sarah.meachem{at}phimr.monash.edu.au)
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Abstract |
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GnRH-immunisation reduced (P<0.001) both type A/ intermediate spermatogonial and type B spermatogonial/ preleptotene spermatocyte number (56% of control) and leptotene/zygotene spermatocyte number (63% of control). Pachytene spermatocyte and round spermatids were reduced to 12% and l% (P<0.01) of control respectively. Testosterone treatment did not increase type A/intermediate spermatogonial number compared with GnRH-immunised controls over the 10-day study period. Treatment with testosterone-esters increased type B spermatogonial/preleptotene spermatocytes and leptotene/zygotene spermatocyte numbers (both being ~83% of control, P<0.05), while T24 cm treatment did not significantly increase their numbers (~73% of control) compared with GnRH-immunised controls. Both treatments increased pachytene spermatocyte and round spermatid numbers to 55% and 8% of control respectively. Co-administration of ICI 182780 had no effect on any of these germ cell numbers. We conclude that oestrogen action plays no role in the short-term restoration of spermatogenesis by testosterone in the GnRH-immunised rat.
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Introduction |
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(ER) (Eddy et al. 1996) and aromatase gene (ArKO) have impaired spermatogenesis (Robertson et al. 1999). The adult ER knock-out testis is grossly dismor-phic, probably due to back pressure of luminal fluids as oestrogen regulates fluid reabsorption in the head of the epididymis (Hess et al. 1997). Normal rats (Oliveira et al. 2001, 2002) and mice (Cho et al. 2003) treated with an ER antagonist (ICI 182780) also provide compelling evidence that testicular disruption is secondary to dilation of the rete testis and efferent ductule lumens due to lack of action by an oestrogen-dependent Na+/H+ exchanger necessary for fluid and electrolyte reabsorption (Zhou et al. 2001). On the other hand, a lack of a direct oestrogenic effect on the seminiferous epithelium is thought to account for the phenotype seen in the ArKO mouse wherein germ cells early in spermiogenesis are the primary site of impairment with no changes being reported in spermato-gonial number (Robertson et al. 1999). Spermatogenesis in the ERß knock-out animal is presumed normal as mice are fertile (Krege et al. 1998), which is surprising given that ERß is present on Sertoli and germ cells, while the combined ERß nock-out animal exhibits a similar phenotype to the ER knock-out mouse (Couse et al. 1999, Dupont et al. 2000). Spermatogenesis is regulated by FSH and testosterone independently and synergistically (McLachlan et al. 2002). In brief, FSH plays a major role in regulating spermato-gonial development in the adult rat (Meachem et al. 1998, 1999). Both FSH and testosterone support spermatocyte maturation, while testosterone is considered essential for spermatid development. In rodent and human models of combined FSH and testosterone deficiency, severe sper-matogenic disruption is observed (McLachlan et al. 2002). For example in the gonadotrophin-releasing hormone (GnRH)-immunised rat, a model that lacks both FSH and LH/testosterone, reductions in spermatogonial and spermatocyte populations to 50% and 10% of control, respectively, are observed while mature spermatids disappear. Exogenous testosterone (e.g. 24 cm Silastic s.c. implant) restores spermatogenesis to near normal by partially restoring testicular testosterone levels (to 10–20% of control) and by the restoration of pituitary FSH by GnRH-independent mechanisms (Meachem et al. 1998, Pratis et al. 2003). This restoration of serum FSH is not always observed (Awoniyi et al. 1989). Whether testosterone-induced spermatogenesis is influenced by a metabolite of testosterone (e.g. oestrogen) is not known. Other data have reported that spermatogenesis is supported by the non-aromatisable androgen dihydrotesto-sterone (DHT), suggesting that aromatisation is not an absolute requirement in gonadotrophin-deficient mice (Singh et al. 1995) and rats (Huang et al. 1987).
In regard to the restoration of spermatogonial development by exogenous testosterone, the data are conflicting. Low-dose testosterone (6 cm Silastic implants) was ineffective in restoring spermatogonial development in GnRH-immunised rats (Meachem et al. 1998), but other studies have shown that low doses of testosterone stimulates spermatogonial number in the hypophysectomised rat (Huang et al. 1987, Sun et al. 1989). On the other hand, an inhibition of spermatogonial development has been suggested in studies using high doses of exogenous testosterone (24 cm Silastic implants) in the GnRH-immunised rat model which may be the reason that spermatogenesis is not quantitatively normal (McLachlan et al. 1994a). Meistrich and colleagues have provided compelling evidence that high testicular testosterone levels inhibited spermatogonial development following exposure to radiation (Meistrich & Kangasniemi 1997, Shuttlesworth et al. 2000) or chemotherapeutic procarbazine in rats (Meistrich et al. 1999), and in the juvenile spermatogonial depletion mutant mouse model (Matsumiya et al. 1999, Shetty & Weng 2004). The mechanism by which testosterone might regulate spermatogonial development in these paradigms is unclear, however it is postulated that a metabolite of testosterone, such as oestrogen, may be involved and this concept is plausible since ERß has been immuno-localised to the rat Sertoli cell and spermatogonia (Saunders et al. 1998).
In order to explore the role of oestrogen in the regulation of spermatogenesis, we examined the restoration of rat germ cell number following a period of regression induced by GnRH-immunisation. In this model, exogenous testosterone treatment partially restores sperm production by both stimulating pituitary FSH secretion and by partially restoring testicular testosterone levels, with the latter also providing substrate for aroma-tisation to oestrogen. The aim of this study was to determine whether the restoration of spermatogenesis by exogenous testosterone was affected by the inhibition of oestrogen action that was achieved by the co-administration of the potent ER antagonist ICI 182780, which targets both ER and ERß (Kuiper et al. 1997, Tremblay et al. 1998, Wakeling 2000). The germ cell response was quantified using the optical disector (sic) stereological technique.
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Materials and Methods |
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Adult Sprague–Dawley rats (75–90 days of age, 350–450 g) were obtained from the Monash Animal House (Clayton, Australia) and maintained at 20 °C in a fixed 12 h light:12 h darkness cycle with free access to food and water. This study was approved by the Monash University Animal Ethics Committee.
Steroid implants
Testosterone (Sigma) Silastic implants (Dow Corning, Corp., Midland, MO, USA) were prepared using medical-grade polydimethylsiloxane tubing (Dow Corning; inner diameter, 1.98 mm; outer diameter, 3.18 mm), and medical adhesive silicone type A as previously described (Robaire et al. 1979). Testosterone implants were 8 cm (3 x 8 cm=24 cm) in length.
Experimental design: spermatogenic suppression
Adult rats were actively immunised with a proprietary GnRH immunogen preparation (BA-1666–4, Biotech, Sydney, Australia) incorporating an adjuvant free of mycobacterial components (Stewart et al. 1992). The immunisation protocol and its effects have been described previously (McLachlan et al. 1994a). Briefly, the GnRH immunogen was administered at a dose of 100 µg protein at a single site every 4 weeks until completion of the study. The control rats received adjuvant only. The response to immunisation was assessed after 12 weeks by measuring testicular volumes by palpation under anaesthesia. A testicular regression to less than 0.55 ml was considered to be an indication of successful immunisation (McLachlan et al. 1994b). All rats showed testicular regression and received a final booster at week 12.
Experimental design: spermatogenic restoration
In order to study the effects of testosterone on the restoration of spermatogenesis in the absence and presence of oestrogen, GnRH-immunised rats (n=7–8 per group) received testosterone Silastic implants (T24 cm) or subcutaneous testosterone esters (25 mg per rat (T25 mg) Sustanon100 (20% testosterone propionate+40% testosterone phenylpropionate+40% testosterone isocaproate); Organon Australia, Sydney) every third day) for 10 days in combination with either the ER antagonist ICI 182780 (2 mg/ml ICI 182780 in oil per kg body weight, s.c. daily injection (Faslodex-Astra-Zeneca, Macclesfield, UK)) or vehicle. ICI 182780 was dissolved in absolute ethanol, and then peanut oil (1:9, v:v) was added prior to evaporation of the solvent under a N2 gas stream. The vehicle was prepared in the same way but the drug was omitted.
This dose of ICI 182780 was chosen based on the following criteria: (i) a 10-fold lower dose compared with that used in this study blocked oestrogen action in human breast cancer patients (Howell et al. 2002) and prevented blastocyst implantation in rats (Dao et al. 1996); (ii) a similar dose to that used in this study disrupted the male reproductive tract of rodents, specifically by inducing dilation of the efferent ductules (Oliveira et al. 2001, 2002, Cho et al. 2003). Two doses of testosterone (T24 cm implant and T25 mg ester) were used to induce graded levels of serum testosterone to test whether higher levels of serum testosterone have detrimental effects on spermato-gonial development.
Tissue collection The left testis was removed (prior to perfusion of the right testis) and frozen in liquid nitrogen and stored at –80 °C prior to assessment of testicular oestrogen levels. As testicular oestrogen concentrations were a priority and tissue mass limited, testicular testosterone levels were not measured but have been previously well characterised in this model and reported elsewhere (McLachlan et al. 1995, Meachem et al. 1998, Pratis et al. 2003). Details of the testis perfusion procedure with Bouin’s fixative, tissue collection, processing, methacrylate embedding, sectioning (25 µm), periodic acid–Schiff’s staining have been previously described (Meachem et al. 1997). Blood was collected by cardiac puncture prior to whole-body perfusion as previously described (Meachem et al. 1997).
Cell number estimates The optical disector method (reviewed in Wreford 1995) was used to determine the total number of cells per testis, as previously described (McLachlan et al. 1994b, Meachem et al. 1999). Germ cells were identified using morphological criteria of Russell and colleagues (Russell et al. 1990) as detailed elsewhere (McLachlan et al. 1994b). A total of between 80 and 160 nuclei of each cell type were counted per rat. A set of unbiased counting frames in each field (with the area of each frame being 1151 µm2) was used to count spermato-gonia and primary spermatocytes, whereas a set of 4 frames (with the area of each frame being 576 µm2) was used to count round and elongated spermatids. Round spermatids were counted in 2 of the 4 counting frames depending on their frequency. As previously determined, no correction for shrinkage was necessary (McLachlan et al. 1995, Meachem et al. 1996). Germ cells were counted in the following categories: type A/intermediate spermatogonia; type B spermatogonia/preleptotene spermatocyte; leptotene/zygotene spermatocyte; pachytene spermatocyte associated with stages I to XIV; round (steps 1 to 8) and elongated spermatids (steps 9 to 19). Germ cells were initially categorised into smaller groupings, but given that no effects were found for testosterone alone or in combination with ICI 182780, germ cells were grouped in broader categories.
Hormone assays
Serum LH was measured by an immunofluorometric assay as previously described (Haavisto et al. 1993). All samples were run in triplicate across one assay. The sensitivity of the assay was 7.8 pg/ml with a within-assay coefficient of variation of 11.0%.
Serum testosterone levels were measured by RIA utilising iodinated histamine–testosterone in combination with an acidic buffer (pH 5.1) to disrupt binding between testosterone and binding proteins in unextracted serum (O’Donnell et al. 1996a). Serum samples were assayed in triplicate across a single assay. Assay sensitivity was 0.6 ng/ml with a within-assay coefficient of variation of 6%.
Serum FSH levels were measured using reagents from the NIDDK reagent program, including iodinated rat FSH (rFSH-8), rat FSH antiserum (rFSH-I7), normal rabbit serum, and rat FSH RP-2 as standard ranging from 0.76–100 ng/ml. Goat anti-rabbit immunoglobulin G (IgG) was used as precipitating second antibody. All samples were assayed in triplicate within the one assay. The sensitivity of the assay was 1.56 ng/ml, with a within-assay coefficient of variation of 14%.
Serum and testicular oestrogen concentrations were determined following extraction on Sep-pak C18 cartridges (Waters, Bedford, MA, USA) in an identical manner as described elsewhere for testosterone extraction (O’Donnell 1996b). Steroid recoveries were monitored throughout by the parallel processing of a group (n=7 animals) of normal control rat sera and testes to which had been added ~10 000 d.p.m. of radiolabelled [3H]oestra-diol (2,4,6,7,16,17-3H(N) oestradiol, 110–170 Ci/mmol, NEN Life Science Products, Boston, MA, USA). These recoveries in the RIA tube were 91.9 ± 6.0 and 73.2± 7.1% (mean± S.D.) for sera and testes respectively. Oestradiol concentrations were then determined by RIA using an in-house iodinated histamine–oestrogen tracer (10 000 c.p.m. per 100 µl in assay buffer (0.1% (w/v) gelatin in 0.1 M PBS (0.154 M NaCl), pH 7.4), 100 µl primary antibody (rabbit anti-oestrogen 4410, Diagnostic Systems Laboratories, Webster, TX, USA) diluted 1:6 in assay buffer, and sample in a final volume of 400 µl. The assay was incubated for 1 h at ambient temperature and then overnight at 4 °C, prior to the addition of second antibody (100 µl goat anti-rabbit IgG diluted 1:20 in assay buffer) for a further incubation at ambient temperature for 30 min. Complexes were precipitated by the addition of 1 ml of 6% polyethylene glycol 6000 for 30 min at 4 °C, after which tubes were centrifuged (30 min, 3000 g). Pellets were then stabilised by the addition of 50 µl of 5% (w/v) potato starch and tubes were re-centrifuged (15 min, 3000 g), drained and counted in a 
-counter. A typical standard curve ranged from 0.08 to 40 pg/100 µl (ED50=5.4 pg/100 µl) with the sensitivity of the assay being 0.8 pg/ml. Samples were assayed in triplicate at multiple dilutions across two to three assays. The within-assay variation was 6.0%, and the between-assay variation was 23%. The specificity of the primary antibody has been described by the manufacturer, which includes cross-reactivities of 2.40, 0.64 and <0.01% for estrone, estriol and testosterone respectively. The cross-reactivity of the ICI 182780 antagonist in this assay was determined to be 0.004%, which at the dosage of antagonist given to the animals would correspond in the RIA to an approximately 40-fold increase in the oestradiol levels actually found in serum. While this calculation does not take into account removal by metabolic clearance, the potential for interference by the ICI antagonist in the RIA was considered too great to assay samples directly from these groups without prior separation.
Statistics
Statistical analyses were performed using Sigmastat for Windows version 2.0 (Jandel Corporation, CA, USA) with an initial assessment of homogeneity of variance for all groups. Homogeneous groups were assessed using one-way ANOVA, followed by the Newman–Keuls post-hoc multiple comparisons test, or in the case of unequal variance, Dunn’s method. The data were expressed as means± S.E.M, with n=4–7 rats per group.
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Results |
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GnRH immunisation reduced testicular weights to 18% of control levels (P<0.001, Table 1
). Testosterone administration increased testicular weights (P<0.001) compared with GnRH-immunised rats (to 35% of control values). Testosterone in combination with ICI 182780 had no effect on testicular weight compared with testosterone alone-treated rats (Table 1).
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Following GnRH immunisation, serum testosterone levels were suppressed to 7% of control values (Table 1
). Serum testosterone levels were increased (P<0.001) ~100-fold and more than 500-fold above GnRH-immunised control levels following treatment with T24 cm Silastic implants and T25 mg ester injections respectively. Administration of ICI 182780 had no effect on serum testosterone levels in T24 cm-treated rats compared with T24 cm alone, however co-administration of ICI 182780 with testosterone ester reduced serum testosterone levels by ~30% (P<0.01).
Serum FSH levels were suppressed by GnRH immu-nisation (P<0.001) to near the detection limit of the assay (Table 1). Following testosterone treatment alone, FSH levels were partially or completely restored in T24 cm- and testosterone ester-treated rats respectively (Table 1). This rise in serum FSH was completely (T24 cm) or partially (testosterone ester) prevented following administration of ICI 182780, therefore showing that ICI 182780 had a biological effect at the pituitary level.
Serum LH was suppressed by GnRH immunisation to undetectable levels and remained so despite all subsequent hormone treatments (Table 1).
Serum oestradiol levels remained unchanged following GnRH immunisation (Table 1). Administration of 24 cm of testosterone did not alter serum oestradiol levels, however administration of testosterone esters increased serum oestradiol levels (P<0.01) 2-fold above control levels.
Testicular oestradiol levels were increased 3.5-fold (P<0.05) following GnRH immunisation compared with control (Table 1). Testicular oestradiol levels tended to be lower following testosterone treatment compared with GnRH-immunised controls, however there was a high within-group variation and no statistical differences were seen.
Germ cell estimates
Suppression phase In response to GnRH immunisation, all germ cell numbers were suppressed (P<0.001) (Fig. 1 and Table 2): type A/intermediate spermatogonia and type B spermatogonia/preleptotene spermatocytes were reduced to 56% of control. Leptotene/zygotene spermatocytes were reduced to 63% of control, while pachytene spermatocytes were suppressed to 12% of control values. Round spermatids were 1% of control values, with only a few step 8 round spermatids observed. Similarly, only a few elongating spermatids were present while no elongated spermatids were seen.
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). Co-administration of ICI 182780 with testosterone had no impact on type A/intermediate spermatogonial number. In contrast, T25 mg increased (P<0.05) type B spermatogonial/ preleptotene spermatocyte number above that of GnRH-immunised rats to ~84% of control values, whereas T24 cm did not (Fig. 1). Co-administration of ICI 182780 with testosterone (T24 cm or T25 mg ester) made no difference to type B spermatogonial/preleptotene spermatocyte number. Both doses of testosterone tended to increase leptotene/zygotene spermatocyte number above that of GnRH-immunised rats to ~85% of control values, although this did not achieve significance (Fig. 1). Again, co-administration of ICI 182780 with testosterone did not alter the leptotene/zygotene spermatocyte number (Fig. 1).
In response to both doses of testosterone treatment, the pachytene spermatocyte number was increased (P<0.001) to a similar extent to 55% of control values (Table 2). The inhibition of oestrogen action did not affect this increase. Both doses of testosterone also increased round spermatid number (P<0.001) to 8% of control values, with co-administration of ICI 182780 again having no significant effect. Elongated spermatid number increased in response to T25 mg (P<0.01) but not following T24 cm treatment. Later spermatids (steps 15 to 19) were almost never seen. ICI 182780 treatment did not significantly affect elongated spermatid number.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To antagonise local oestrogen action in the testis, this study employed a similar dose (g/kg body weight) of ICI 182780 to that reported elsewhere in rats to cause signifi-cant changes to both testis weight and dilation of seminiferous tubules after 52 days of treatment, and severe atrophy by day 150 (Oliveira et al. 2001). These testicular effects were probably a result of an accumulation of fluid following inhibition of reabsorption in the efferent duc-tules (Oliveira et al. 2001), similar to the phenotype seen in the ER-deficient mouse (Eddy et al. 1996). In a follow-up study these authors confirmed that changes in the efferent ductules preceded those in the testes indicating that the testicular effects of ICI 182780-treated rats were secondary to the epididymal effects (Oliveira et al. 2002). These data confirm that the antagonist doses used in our study would be expected to substantially impair ER-mediated effects, consistent with the observation of a suppression of serum FSH in this study.
Germ cell regulation
The GnRH-immunised rat model displays well-characterised reductions in all germ cell types (McLachlan et al. 1995, Meachem et al. 1998) but especially developmental arrest in mid-spermiogenesis (McLachlan et al. 1994a). Administration of testosterone (doses of 6 and 24 cm Silastic implants) to GnRH-immunised rats promptly restores serum FSH by a GnRH-independent mechanism and hence this model is one of acute testosterone and FSH restoration (McLachlan et al. 1994a, Meachem et al. 1998). However, for reasons that are unclear, this increase in serum FSH has not always been reported by others in testosterone-treated GnRH-immunised rats (Awoniyi et al. 1989).
In regard to the hormonal regulation of spermatogonial number, in the GnRH-immunised rat model FSH is required for their restoration (McLachlan et al. 1995, Meachem et al. 1998) with low testosterone (6 cm) being ineffective in restoring spermatogonial number in the absence of FSH (Meachem et al. 1998). High-dose serum testosterone (5-fold normal as induced by T24 cm (Meachem et al. 1998)), independent of FSH action, fails to increase spermatogonial number above that of the GnRH-immunised rat (Ebling et al. 2000). High-dose testicular testosterone (4-fold normal) observed after irradiation in rats has been found elsewhere to inhibit spermatogonial differentiation, and lowering these levels, even transiently, was sufficient to enhance the recovery of spermatogonia (Ogawa et al. 1999). Therefore it was postulated that metabolism of testosterone to oestrogen by aromatisation may impair spermatogonial development (Meistrich & Kangasniemi 1997). This explanation seems plausible considering that ERß has been immuno-localised to rat Sertoli cells and spermatogonia (Saunders et al. 1998). However, our data demonstrate that treatment of rats with ICI 182780 did not ameliorate the proposed inhibitory effects of high levels of testosterone on spermato-gonial numbers, suggesting that it is not oestrogenic in nature. Alternative mechanisms for the inhibitory effects of high-dose testosterone on spermatogonial number could include: (i) that testosterone itself acts in this fashion in these specialised settings; (ii) that testosterone treatment results in the production of an additional product arising from elsewhere (e.g. in the liver); or (iii) another metabo-lite of testosterone, such as its 5-alpha-reduced product DHT, may be involved, noting that the administration of testosterone to GnRH-immunised rats elevated testicular DHT levels (Pratis et al. 2003).
Evidence supporting an oestrogenic effect(s) on spermato-gonial development is lacking. Consistent with this, spermatogonial numbers are normal in mice lacking a functional aromatase gene (Robertson et al. 1999) suggesting that oestrogen plays no role in this model. Transplantation of ER-deficient germ cells into wild-type testes resulted in qualitatively normal spermatogenesis (Mahato et al. 2000). However, an ERß-directed mechanism is plausible given that ERß receptors are present on spermato-gonia (Saunders et al. 1998), but no similar transplantation experiment has been reported with ERß-deficient germ cells.
Our data show that testosterone supports type B spermato-gonial and primary spermatocyte development and that this effect is independent of oestrogen action as it was unaffected by ICI 182780 treatment. Furthermore, the action of testosterone presumably involves enhanced cell survival or mitosis of type B spermatogonia rather than increased precursor cell entry since type A spermatogonial number was unaffected. Other reports suggest that testosterone supports type B spermatogonial and primary spermatocyte number independent of type A spermatogonia (Meachem et al. 1997, El Shennawy et al. 1998, Franca et al. 1998, Russell et al. 1998) and that testosterone exerts its effects by preventing their degeneration (El Shennawy et al. 1998, Russell et al. 1998).
Exogenous testosterone support for meiosis and early spermiogenesis has been widely reported in rats (Muffly et al. 1993, 1994, McLachlan et al. 1994b, O’Donnell et al. 1996a, Saito et al. 2000) and genetically modified mice models (Chang et al. 2004, De Gendt et al. 2004, Spaliviero et al. 2004). Oestrogen receptor inhibition in this study had no effect on the restoration of spermatocyte and spermatid number by testosterone. This is consistent with other studies (Chen et al. 1994, Singh et al. 1995) suggesting that aromatisation is not required to support spermatogenesis; however it has been shown elsewhere that early spermiogenesis is sensitive to oestrogen action (Robertson et al. 1999). Although testicular testosterone was not measured in the current study due to the limited amount of available tissue, our previous reports have consistently shown that T24 cm treatment of GnRH-immunised rats elevates total testicular testosterone levels to ~10% of control levels (Meachem et al. 1998, Pratis et al. 2003), compared with 3% in the GnRH-immunised control. Using the same rat model, others have demonstrated an elevation in interstitial fluid testosterone to ~20% of control following T24 cm treatment (Awoniyi et al. 1989, 1992), with this variation most likely to be due to the different assay systems and tissues measured. While the effect of T25 mg esters on testicular testosterone has not previously been determined in this model, it is considered that this treatment would elevate testosterone to ~20% of control levels; this assumption is based on data where the same treatment was administered to rats selectively withdrawn of testicular testosterone by low-dose testosterone and oestrogen implants, where testicular testosterone also falls to 3% of control (Meachem et al. 1997). Numerous studies (Boccabella 1963, Cunningham & Huckins 1979, Marshall et al. 1984) have reported that spermatogenesis can be maintained with 10–20% of normal testicular testosterone.
This study also highlights that germ cell types have differing dependencies for testosterone. Support of type B spermatogonia (El Shennawy et al. 1998, Meachem et al. 1998), spermatocytes (McLachlan et al. 1994b, Meachem et al. 1997, 1998) and spermatid maturation (McLachlan et al. 1994b, Meachem et al. 1997, 1998, O’Donnell et al. 1996a,b, 1999) requires testicular testosterone, albeit at much lower levels (~10–20% of control) than the normal setting produces (Boccabella 1963, Cunningham & Huckins 1979, Marshall et al. 1984). In contrast, testosterone provides no support for the restoration of type A spermatogonial number (Meachem et al. 1997, 1998), while supraphysiological levels of serum (Meachem et al. 1997, 1998) and testicular testosterone (Meistrich & Kangasniemi 1997, Matsumiya et al. 1999, Meistrich et al. 1999, Shuttlesworth et al. 2000, Shetty & Weng 2004) have a detrimental effect on type A spermatogonial development. The mechanism(s) behind this differential response is not known.
Hypothalamo-pituitary–testicular axis regulation
Serum FSH levels were significantly reduced by concomitant ICI 182780 administration providing support for the notion that oestrogen acts in a positive manner on the male pituitary. Others have noted a stimulatory effect of low-dose oestrogen on FSH in neonatal (Tena-Sempere et al. 2000) and hpg mice (Ebling et al. 2000). In the setting where the hypothalamic-pituitary axis is intact, it is clear that oestrogen exerts an inhibitory effect on gonadotrophin secretion (de Jong et al. 1975, reviewed in O’Donnell et al. 2001). GnRH facilitates FSH secretion directly from the pituitary by augmenting the GnRH receptor signal trans-duction pathway, with oestrogen also being able to facilitate this pathway and stimulate FSH secretion (reviewed in Shupnik 1996). Presumably in our study, in the absence of effective GnRH action on the gonadotrophs, oestrogen (via metabolism from exogenous testosterone) promotes the release of FSH and the ER antagonist inhibits this effect. Taken together these data indicate that oestrogen can participate in both negative and positive effects on the pituitary in the male.
Oestradiol concentrations
Studies describing oestrogen concentrations in the male reproductive tract are reviewed elsewhere (Hess 2000). Rat serum oestradiol concentrations in this study are similar to those reported by de Jong and colleagues (de Jong et al. 1973). While rat whole testicular oestradiol concentrations have not previously been described, oestra-diol concentrations in the testicular vein (de Jong et al. 1973), seminiferous tubules (de Jong et al. 1974) and rete testis (Free & Jaffe 1979) have been reported. Our testicular oestradiol data are in agreement with levels reported in dissected seminiferous tubules (de Jong et al. 1974), with concentrations within the seminiferous epithelium being nine times higher than in serum (de Jong et al. 1974). In this study testicular oestradiol levels, regardless of hormonal manipulation, were ~10-fold higher than circulating levels, consistent with de Jong and colleagues (de Jong et al. 1973) and data from a variety of species (reviewed in de Jong et al. 1973).
Unexpectedly, serum concentrations of oestradiol in GnRH-immunised rats remained at control levels, while oestradiol concentrations in the testis were elevated 3-fold above control values, even in the presence of very low levels of testicular testosterone (3% of control, (Meachem et al. 1998, McLachlan et al. 1994a)). Testicular oestradiol concentrations may be elevated in this gonadotrophin-depleted model for one of several reasons. (1) Oestrogen production in the rat may be, at least in part, GnRH independent. Other tissues such as adipose tissue (pre-adipocytes and stromal mesenchymal cells) and bone (osteoblasts) produce oestrogen and are not under GnRH control (reviewed in Simpson et al. 2002). Cytokines, such as tumour necrosis factor (TNF)- and interleukin (IL)-11, regulate oestrogen production in these tissues (reviewed in Simpson et al. 2002). As TNF- and IL have been described in the testis, these cytokines may play a local role (reviewed in Gnessi et al. 1997). It has been shown that low levels of testicular testosterone are constitutively present in gonadotrophin-depleted mice suggesting a GnRH-independent mode of testosterone production (Zhang et al. 2003), which may also be true for oestrogen. (2) Testosterone may negatively regulate aromatase activity, thus aromatase activity may be up-regulated in the GnRH-immunised model resulting in an increase in oestrogen levels. Presumably only Leydig and/or Sertoli cell aromatase activity would contribute to this effect, since germ cell loss is extensive in GnRH-immunised animals. (3) It may be possible that oestrogen metabolism was altered in the GnRH-immunised rat and as a consequence oestradiol accumulated. Serum and testicular oestradiol concentrations were not determined in the ICI 182780 antagonist-treated rats in this study because the antagonist cross-reacted in the oestradiol RIA.
It is concluded that oestrogen action plays no role in the short-term restoration of spermatogenesis by testosterone in the GnRH-immunised rat. More particularly, the failure of high-dose testosterone to fully restore type A spermatogonia is not ameliorated by inhibition of ER action, and no evidence was found for the modulation of later germ cell progression by oestrogen. This study also provides new data regarding oestradiol concentrations in gonadotrophin-depleted rats, with an unexplained increase in testicular oestradiol concentrations in this model. In addition, this study provides further evidence that oestro-gen can participate in the positive regulation of pituitary FSH in the rat in the absence of GnRH bioactivity.
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Acknowledgements |
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Funding |
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This study was supported by the Lalor Foundation, USA, Wellcome Trust Fellow Scheme, UK (grant 058479 to S M) and the National Health and Medical Research Council of Australia (grant 241000 to S M, D M R, R M , P S). The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.
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References |
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Received 4 March 2005
Accepted 9 March 2005
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