Androgens and gonadotrophins play crucial roles in the development and function of the male reproductive tract, and spermatogenesis. In addition, the requirement of estrogens for male fertility has been established recently in experiments, in which aromatase- and estrogen receptor-knockout mice (ArKO and ERKO) were used. The testicular phenotype of these mice was characterized by seminiferous tubule atrophy and several abnormalities in sperm morphology. In ArKO mice, progressive disruption of spermatogenesis caused by abnormal germ cell development has been described whereas the infertility of the ERKO has been shown to be a result of impaired fluid reabsorption in the efferent ductules. In humans, it has been shown that aromatase deficiency as well as lack of ER expression due to naturally occurring mutations lead to infertility and decreased sperm viability.

Although, a direct effect of estrogens on male reproduction is not yet completely understood, the data obtained in rodents clearly show that estrogens are important to maintain reproductive functions in the male.

In various species, including humans, aromatase, the enzyme responsible for estrogens formation, has been found to be expressed in either Leydig cells or in both compartments of the testis, the interstitial area and seminiferous tubules. In tubules, aromatase has been found in meiotic and post-meiotic germ cells, predominantly in spermatids. Both aromatase expression and estrogen synthesis in testis as well as the exact cellular sites have been extensively reviewed recently. It should be stressed, however, that already more than 20 years ago estrogen effects have been described on morphology of human Leydig cells and testicular steroidogenesis in the rat, but the recent discovery of aromatase and estrogen receptors in male germ cells makes the role of estrogens in spermatogenesis especially interesting.

We suggest that seasonally breeding animals such as bank voles are a useful model for studies of male reproduction since the recrudescence of spermatogenesis is physiologically regulated. This assumption is based on earlier observations that season and length of the light cycle affect reproduction effectiveness, regulate changes in hormonal activity and fertility parameters of sperm. Bank voles, kept under different light cycle conditions in the laboratory, behave similarly as wild animals. They show different phases of reproduction, active or regressive, that mimick spring or autumn photoperiods. Low activity of enzymes involved in steroidogenesis and weak expression of androgen receptors immunoreactivity in male and female voles that were kept under short light cycles have been described previously. In this context, the question arises whether estradiol supplementation or deprivation can induce and/or disrupt spermatogenesis in male immature bank voles bred under short or long photoperiods.

Bank voles (Clethrionomys glareolus, Schreber) were obtained from our own colony (Laboratory of Endocrinology and Tissue Culture, Institute of Zoology, Jagiellonian University, Krakow, Poland) and since 10 generations, they have been reared under short light cycles (6L:18D) or long light cycles (18L:6D). The animals were housed at 18±2°C in polyethylene cages (42×27×18 cm³) furnished with sawdust and wood shavings for bedding. A standard pelleted diet supplemented with seeds of wheat or oat, red beet, apples, and water were provided ad libitum. Fifty immature bank voles (15–20-days old) were injected intraperitoneally (ip), twice a week during 2 weeks, either with 17?-estradiol dissolved in 20 ?l sesame oil (Sigma, St. Louis, MO, USA) at a dose of 0.1 and 10 ?g/g body weight or with the anti-estrogen ICI 182,780 (Faslodex, a gift from Astra Zeneca, Cheshire, UK) at a dose of 10 and 100 ?g/g body weight dissolved in the same solvent. A control group (6 animals) was treated with 20 ?l sesame oil only. These doses were selected on the basis of results of a preliminary dose-dependency study. Irrespective of the photoperiod, the low doses of estradiol (0.01 ?g/g body weight) and ICI (0.1 and 1 ?g/g body weight) did not cause any alterations in tubular structure when compared to controls, whereas higher doses of estradiol and ICI (100 and 1000 ?g/g body weight), respectively, were harmful for the males. The animals were killed by cervical dislocation and the testes were immediately excised. Bouin’s fixative or neutral-buffered formaldehyde were used for routine histological analysis or immunohistochemical staining.

Experiments were performed in accordance with Polish legal requirements, under the licence given by the Commission of Bioethics at the Jagiellonian University, Krakow, Poland.

For morphological analysis, sections of testes from both control and experimental animals were routinely stained with haematoxylin and eosin.

Immunohistochemistry and TUNEL staining was performed on deparaffinized and rehydrated 6-?m-thick paraplast sections (Monoject Scientific Division of Scherwood Medical, St. Louis MO, USA). To optimize immunohistochemical staining, sections were immersed for 2×5 min in 10 mM citrate buffer (pH 6.0) at high temperature to unmask antigen. After heating in the microwave, the container holding the sections was removed and allowed to cool for 15 min at room temperature. Then, sections were washed and the immunostaining procedure started. Nonspecific staining was blocked twice, first with 3% H₂O₂ in methanol for 15 min to inhibit endogenous peroxidase activity, and second with 10% normal goat serum for 30 min at room temperature to block nonspecific binding sites. After that, sections were processed for visualization of aromatase using the immunohistochemical technique described elsewhere. Sections were incubated overnight at 4°C in a humidified chamber in the presence of the primary antibody, a rabbit polyclonal antibody against human placental P450 aromatase (dilution, 1:400; R-10-2; a generous gift from Dr Yoshio Osawa, Hauptman-Woodward Medical Research Institute, Buffalo NY, USA). This antibody was raised in a rabbit against human placental P450 aromatase that had been purified previously by immunoaffinity using a monoclonal antibody against P450 aromatase. It was developed with the support of US Public Health Service Research Grant # HD P4945 from NICHHD. Next, biotinylated secondary antibody, goat anti-rabbit IgG (dilution, 1:400; Vector, Burlingame, CA, USA) was applied. Finally, avidin–biotinylated horseradish peroxidase complex (ABC/HRP; dilution, 1:100; Dako, Glostrup, Denmark) was used. After each step in these procedures, sections were carefully rinsed with Tris-buffered saline (TBS; 0.05 M Tris-HCl and 0.15 M NaCl, pH 7.6); all antibodies were also diluted in TBS buffer. Peroxidase activity was visualized using 0.01% H₂O₂, 0.05% diaminobenzidine, and 0.07% imidazole (Sigma) dissolved in TBS. Then, sections were rinsed in tap water. Experiments were repeated several times and in each staining experiment, sections were incubated in the presence of irrelevant IgG instead of the primary antibody as negative control.

Deparaffinized and rehydrated paraplast sections from both control and experimental groups were also stained using terminal deoxyribonuclotidyl transferase-mediated dUTP nick end labeling (TUNEL) method. Apoptotic cells were visualized using the in situ death detection kit POD (Roche Molecular Biochemicals, Mannheim, Germany). Peroxidase activity was visualized using the 3,3? diaminobenzidine tetrachloride liquid dropper system (Sigma). As negative controls, sections were processed without terminal deoxynucleotidyltransferase (TdT) buffer.

Finally, sections were examined with a Leica DMR microscope (Wetzlar, Germany) using bright field illumination and/or Nomarski interference contrast microscopy.