17β-Estradiol activates GPER- and ESR1-dependent pathways inducing apoptosis in GC-2 cells, a mouse spermatocyte-derived cell line

Molecular and Cellular Endocrinology 355 (2012) 49–59

Contents lists available at SciVerse ScienceDirect

Molecular and Cellular Endocrinology journal homepage: www.elsevier.com/locate/mce

17b-Estradiol activates GPER- and ESR1-dependent pathways inducing apoptosis in GC-2 cells, a mouse spermatocyte-derived cell line Adele Chimento a,1, Rosa Sirianni a,1, Ivan Casaburi a, Carmen Ruggiero a, Marcello Maggiolini a, Sebastiano Andò b, Vincenzo Pezzi a,⇑ a b

Department of Pharmaco-Biology, University of Calabria, 87036 Arcavacata di Rende, Cosenza, Italy Department of Cell Biology, University of Calabria, 87036 Arcavacata di Rende, Cosenza, Italy

a r t i c l e

i n f o

Article history: Received 29 July 2011 Received in revised form 17 January 2012 Accepted 19 January 2012 Available online 27 January 2012 Keywords: GPER ESR1 Spermatocytes Estrogen Apoptosis

a b s t r a c t In mammals, spontaneous apoptosis is observed particularly in differentiating spermatogonia and in spermatocytes. 17b-Estradiol (E2) in primary rat pachytene spermatocytes (PS) binds estrogen receptor a (ESR1) and GPER to activate EGFR/ERK/c-Jun pathway leading to up regulation of proapoptotic factor bax. Aim of this study was to clarify the effector pathway(s) controlling spermatocytes apoptosis using as model GC-2 cells, an immortalized mouse pachytene spermatocyte-derived cell line, which reproduces primary cells responses to E2. In fact, in GC-2 cells we observed that ESR1 and GPER activation caused rapid ERK and c-Jun phosphorylation, bax up-regulation, events associated with apoptosis. We further investigated the apoptotic mechanism demonstrating that E2, as well as ESR1 and GPER specific agonists, induced sustained ERK, c-Jun and p38 phosphorylation, Cytochrome c release, caspase 3 and endogenous substrate Poly (ADP-ribose) polymerase (PARP) activation and increased expression of cell cycle inhibitor p21. When ESR1 or GPER expression was silenced, E2 was still able to decrease cell proliferation, only the concomitant silencing abolished E2 effect. These results indicate that GC-2 cells are a valid cell model to study E2-dependent apoptosis in spermatocytes and show that E2, activating both ESR1 and GPER, is able to induce an ERK1/2, c-Jun and p38-dependent mitochondrion apoptotic pathway in this cell type. Ó 2012 Elsevier Ireland Ltd. All rights reserved.

1. Introduction It is well known that normal testicular development and maintenance of spermatogenesis are controlled by gonadotrophins and testosterone, whose effects are modulated by a complex network of locally produced factors, including estrogens (Carreau et al., 2003). Indeed, the androgen/estrogen balance is under a fine tuning via endocrine and paracrine factors, but is also dependent on aromatase, an enzymatic complex localized in most of the testicular cells responsible for androgens conversion into estrogens (Carreau et al., 2009). Despite the acknowledgment of estrogens as part of spermatogenesis regulating factors, the mechanisms by which they influence spermatogenesis remain uncertain. In order to exert a biological role, testicular estrogens should interact with estrogen receptors a (ESR1) and b (ESR2), which in turn modulate transcription of specific genes. Some studies in vivo and in vitro revealed that estrogen can act as germ cell survival factor and that this effect is dose-dependent (Pentikäinen et al., 2000). For example, estradiol (E2) prevents apoptosis of germ ⇑ Corresponding author. Tel.: +39 0984 493157; fax: +39 0984 493271. 1

E-mail address: [email protected] (V. Pezzi). These authors contributed equally to this work.

0303-7207/$ - see front matter Ó 2012 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.mce.2012.01.017

cells within human seminiferous tubules in vitro even in the absence of gonadotropins (Pentikäinen et al., 2000). However proapoptotic effects of E2 on spermatogenesis have also been observed. Mishra and Shaha (2005) demonstrated that estradiol-induced apoptosis in different germ cells occurred in vitro in the absence of somatic cells, demonstrating independent (not mediated by Sertoli cells) capability of germ cells to respond to estrogen activating apoptosis. However, it has been reported that, in addition to the genomic actions, estrogens can trigger rapid signaling responses that are initiated at the plasma membrane and mediated through activation of intracellular signaling cascades independent of nuclear translocation (Levin, 2008). Recently, an orphan G protein-coupled receptor (GPER) with high-affinity and low-capacity estrogens binding properties was identified at both the plasma membrane (Funakoshi et al., 2006) and the endoplasmic reticulum (Revankar et al., 2005). GPER was cloned from several mammalian species and shown to be expressed in a variety of human and rodent estrogen target tissues (Brailoiu et al., 2007; Chagin and Savendahl, 2007; Sakamoto et al., 2007; Wang et al., 2007) including the testis (Isensee et al., 2009). We recently demonstrated expression of GPER in mouse spermatogonia cell line (Sirianni et al., 2008), rat pachytene spermatocytes (PS) (Chimento et al., 2010) and round spermatids (RS) (Chimento et al., 2011).

50

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

Studies performed on human (Pentikäinen et al., 2000) rat (Bois et al., 2010; D’Souza et al., 2005; Mishra and Shaha, 2005), mouse (Delbes et al., 2004; Ebling et al., 2000) and bank vole (Gancarczyk et al., 2004) germ cells showed that estrogens can determine different effects on different species and/or germ cell types. This phenomenon could be explained by different patterns of expression and/or action of ERs among different species and different germ cell types. Our recent data, demonstrated that E2, in mouse spermatogonial GC-1 cell line, through a cross talk between GPER and ESR1, activates the rapid EGFR/ERK/fos pathway, which in turn stimulates cell proliferation (Sirianni et al., 2008). However in primary rat PS E2, working through both ESR1 and GPER, activates the rapid EGFR/ERK/c-Jun pathway, which in turn induces expression of genes involved in cell apoptosis (Chimento et al., 2010). Moreover, in primary rat RS E2 via ESR1 and GPER, activates pathways involved in the regulation of genes controlling RS apoptosis and differentiation, such as cyclin B1 and bax (Chimento et al., 2011). Interestingly Lucas and colleagues (Lucas et al., 2010) have shown in cultures of immature rat Sertoli cells that the regulatory mechanisms that control cell proliferation (activation of ERs) and antiapoptotic effect (activation of GPER) are remarkably overlapping and involve activation of EGFR-ERK1/2. PS are diploid male germ cells that undergo two meiotic divisions and become haploid RS, which eventually transform into mature elongated spermatozoa. PS represent a very interesting type of spermatogenic cells since before further differentiation they can proliferate or die through apoptosis, they represent the major spermatogenic cell type affected by this phenomenon (Billig et al., 1995; Perrard and Durand, 2009) and a good target to control spermatogenesis. All the studies investigating apoptotic mechanisms activated in germ cells are limited by the possibility to keep these cells in culture only for short times. The availability of an immortalized spermatocyte-derived cell line with the same behavior of primary cultures but easily available for transfection experiments by conventional methods, would be useful to define E2 signaling pathways eliciting apoptosis and to test in vitro drugs that can control cell divisions. In the present study we wanted to test if GC-2 cells (Hofmann et al., 1994) can reproduce the same effects that we observed in cultures of primary PS (Chimento et al., 2010) and further characterize the mechanisms involved in cell apoptosis. In particular in this study we: (i) evaluated estrogen receptor expression; (ii) determined if E2 could trigger apoptosis; (iii) defined the estrogen receptor subtype(s) involved in this mechanism; (iv) defined the signaling pathway activated by E2 to elicit apoptotic effects in GC-2 cells.

2. Materials and methods 2.1. Cell cultures and treatments GC-2 cells (a mouse spermatocyte-derived cell line, obtained from American Type Culture Collection (ATCC) (Manassas, VA, USA) were cultured in Dulbecco’s Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F-12) with phenol red supplemented with 10% fetal bovine serum (FBS), 1% glutamine and 1% penicillin/streptomycin (Sigma–Aldrich, Milano, Italy) (complete medium). Cells were cultured at 37 °C for no more than 10 passages; for experiments cells were plated in complete medium, 24 h later treated in DMEM/F-12 medium without phenol red and serum (serum-free medium) for the indicated times, G1 and 4,40 ,400 -(4-propyl-[1H]pyrazole-1,3,5-triyl) trisphenol (PPT) (Tocris Bioscience, Ellisville, Missouri, USA) and E2 (Sigma–Aldrich) at the indicated concentrations.

TM3 cells, an immature mouse Leydig cell line (ATCC), were cultured in DMEM/F-12 medium supplemented with 5% horse serum (HS), 2.5% FBS, 1% glutamine and 1% penicillin/streptomycin (Sigma–Aldrich). GC-1 cells, a mouse spermatogonia type B cell line (ATCC), were cultured in DMEM/F-12 medium supplemented with 10% FBS, 1% glutamine and 1% penicillin/streptomycin (Sigma– Aldrich). 2.2. RNA extraction, cDNA synthesis and PCR reaction Cells were cultured in complete medium for 24 h in 60 mm dishes (1  106 cells). RNA was extracted using the TRizol RNA isolation system (Invitrogen, Carlsbad, CA, USA). Each RNA sample was treated with DNase I (Ambion, Austin, TX, USA), and purity and integrity of RNA samples was confirmed spectroscopically and by gel electrophoresis prior to use. One lg of total RNA was reverse transcribed in a final volume of 30 ll using the ImPromII Reverse transcription system kit (Promega Italia S.r.l., Milano, Italy). cDNAs were directly used for PCR or diluted 1:3 in nuclease free water to be used for Real time PCR. Samples were aliquoted and stored at 20 °C. Two microliters of cDNAs were used for PCR reactions with gene-specific primers. For ESR1, ESR2 and GPER PCR conditions and primer sequences were previously published (Sirianni et al., 2008). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as housekeeping gene (Sirianni et al., 2008). PCR products were analyzed on a 1% agarose gel and visualized by ethidium bromide staining. 2.3. Western blot analysis Cells were cultured in complete medium for 24 h in 60 mm dishes (1  106 cells), before being treated in serum-free medium. Methods for protein extraction and blots preparation have been previously published (Sirianni et al., 2007). Blots were incubated overnight at 4 °C with (a) anti-GPER polyclonal antibody (1:1000) (MBL International Corporation, Woburn, MA, USA), (b) anti-ESR1 (F-10) antibody (1:500) (Santa Cruz Biotechnology, Santa Cruz, CA, USA), (c) anti-ESR2 (H-150) antibody (1:1000) (Santa Cruz), (d) anti-pERK antibody (1:1000) (Cell Signaling Technology, Beverly, MA, USA), (e) anti-ERK antibody (1:1000) (Cell Signaling Technology), (g) anti-phospho c-Jun (Ser 73) antibody (1:1000) (Cell Signaling Technology), (f) anti-c-Jun antibody (1:1000) (Santa Cruz Biotechnology) (h) anti-phospho p38 MAPK antibody (1:1000) (Epitomics, Inc., Burlingame, CA, USA), (i) anti-p38 MAPK antibody (1:1000) (Cell Signaling Technology), (j) anti-cytochrome c antibody (1:1000) (Santa Cruz Biotechnology), (k) anti-bax antibody (1:1000) (Santa Cruz Biotechnology), (l) anti-bcl-2 antibody (1:1000) (Santa Cruz Biotechnology), (m) anti-caspase 3 antibody (1:1000) (Santa Cruz Biotechnology), (n) anti-caspase 9 antibody (1:500) (Cell Signaling Technology), (o) anti-PARP antibody (1:3000) (Santa Cruz Biotechnology), (p) anti-cyclin D1 (1:1000) (Santa Cruz Biotechnology), (q) anti-cyclin B1 (1:1000) (Santa Cruz Biotechnology), (r) antip21 (1:250) (Santa Cruz Biotechnology). Membranes were incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (Amersham Pharmacia Biotech, Piscataway, NJ) and immunoreactive bands were visualized with the ECL Western blotting detection system (Amersham Pharmacia Biotech). To assure equal loading of proteins, membranes were stripped and incubated overnight with an anti-GAPDH antibody (1:3000) (Santa Cruz Biotechnology). 2.4. Immunofluorescent staining Cells were cultured in complete medium for 24 h on microscope slides (1  105 cells) and then treated in serum-free medium for 6 h for pERK staining. Cells were then fixed using

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

paraformaldehyde (4%) (Sigma–Aldrich) for 30 min at room temperature. Before incubation slides were washed three times for 5 min with phosphate buffer saline (PBS) (Sigma–Aldrich) and then soaked for 3 min with 0.3% of Triton X-100 in PBS. After three washes in PBS, slides were incubated in PBS containing 3% bovine serum albumin (BSA) for 30 min at room temperature and then with a rabbit anti-GPER antibody (1:50) or a rabbit anti-pERK antibody (1:1000) in PBS containing 0.1% BSA (Sigma–Aldrich) for 48 h (for GPER) or over night (for pERK) at 4 °C. On the following day, slides were washed with PBS and then incubated with a FITC-conjugated anti-rabbit immunoglobulin diluted in PBS (Sigma–Aldrich) (1:50 for GPER, 1:500 for pERK) for 1 h at room temperature. After being washed with PBS, slides were incubated with a 1% propidium iodide solution (Sigma–Aldrich) for 30 min or 2-(4-amidinophenyl)-6-indolecarbamidine dihydrochloride (DAPI) (0, 2 lg/ml) (Sigma–Aldrich) for 10 min at room temperature and then analyzed on a fluorescent microscope. Control staining was performed on adjacent serial slides and consisted in replacing the primary antibody with 0.1% BSA in PBS. 2.5. Assessment of cell proliferation Cells were cultured in complete medium in 24 well plates (2  105 cells/well) for 24 h, then treated in serum-free medium for 48 h. Control cells were treated with the same amount of vehicle alone (dimethylsulfoxide), which never exceeded the concentration of 0.01% (vol/vol). [3H]thymidine incorporation was evaluated after a 6 h incubation period with 1 lCi [3H]thymidine per well (Perkin–Elmer Life Sciences, Boston, MA, USA). Cells were washed once with 10% trichloroacetic acid, twice with 5% trichloroacetic acid, and lysed in 1 ml 0.1 N NaOH at 37 °C for 30 min. The total suspension was added to 5 ml optifluor fluid and radioactivity determined in a b-counter. 2.6. Apoptosis detection with Annexin V-FITC Cells were cultured in complete medium for 24 h on microscope slides (1  105 cells), then treated in serum-free medium for 12 h. Annexin V Apoptosis Detection Kit (Santa Cruz Biotechnology) was used for labeling cells undergoing apoptosis. After treatments, cells were washed with 1X Assay Buffer and then incubated with Annexin V-FITC conjugated (0, 5 lg/100 ll) for 15 min. Cells were visualized under a fluorescence microscope at 400 magnification. 2.7. Determination of nuclear morphological changes Cells were cultured in complete medium for 24 h on microscope slides (1  105 cells), then treated in serum-free medium for 24 h. Cells were washed with PBS and fixed in 4% formaldehyde for 10 min at room temperature. Fixed cells were washed with PBS and incubated with 2-(4-amidinophenyl)-6-indolecarbamidine dihydrochloride (DAPI) (0, 2 lg/mL) for 10 min in a humidified chamber, protected from light, at 37 °C. Cells were then washed three times with cold PBS and one drop of mounting solution was added. Cell nuclei were observed and imaged by an inverted fluorescence microscope (400 magnification) with excitation at 350 nm and emission at 460 nm. The number of apoptotic nuclei was determined in at least six randomly selected areas from three cover slips of each experimental group. 2.8. Determination of DNA fragmentation Cells were cultured in complete medium in 100 mm dishes (1  106 cells) for 24 h, then treated in serum free medium for 24 h. To determine the occurrence of DNA fragmentation, total DNA was extracted from control and E2, G1 and PPT treated cells.

51

In brief, the attached and detached cells floating in the medium were collected by scraping and centrifuging (1500 rpm for 5 min at 4 °C). Pellets were washed three times with PBS and then resuspended in DNA laddering lysis buffer (10% NP40, 200 mM EDTA, 0.2 M Tris–HCl pH 7.5). Lysates were centrifuged at 3000 rpm for 5 min at 4 °C. The recovered DNA was incubated with RNase A (final 5 lg/ll) in 1% SDS for 2 h at 56 °C. After addition of proteinase K (final 2.5 lg/ll) samples were incubated for an additional 3 h at 37 °C. DNA precipitation was performed using ethanol/ammonium acetate precipitation O/N at 80 °C. The following day samples were centrifuged at 12,000 rpm for 20 min at 4 °C and washed with 80% ice-cold ethanol. DNA pellets were resuspended in nuclease-free water. Equal amounts of DNA were analyzed by electrophoresis on a 2% agarose gel stained with ethidium bromide (Sigma–Aldrich). 2.9. Comet assay Cells were cultured in complete medium for 24 h in 60 mm dishes (1  106 cells), then treated in serum free medium for 24 h. Comet assay was performed as previously described (Lee et al., 2003; Tice et al., 2000). Briefly, cells were trypsinized with 0.005% trypsin and diluted with equal amount of complete medium. Glass slides were soaked in methanol and frosted by flaming, then coated with 100 ll of 0.8% agarose and covered with coverslips and left at 4 °C overnight. Coverslips were gently removed and a mixture of 30 ll of cell suspension and 50 ll of 0.5% agarose was added rapidly and evenly to the frosted slides and allowed to solidify for 10 min at 4 °C in the dark. Slides were then placed in a tank with lysis solution (2.5 M NaCl, 0.1 M EDTA, 10 mM Tris, 300 mM NaOH, 10% dimethyl sulfoxide, and 1% Triton X-100, pH 10.0) at 4 °C for 1 h in the dark, washed three times with PBS and incubated in fresh alkaline buffer (300 mM NaOH/1 mM EDTA, pH > 13) for 30 min at room temperature to allow the DNA to unwind. Electrophoresis was performed at room temperature in ice-cold alkaline electrophoresis buffer for 35 min at 25 V. After electrophoresis, slides were gently washed with PBS and incubated in neutralization buffer (Tris 400 mM, pH 7.5), let sit for 10 min, stained with 30 ll of DAPI solutions (0, 2 lg/mL), rinsed carefully with PBS and then covered with coverslips. Slides were visualized using a 100 objective on a fluorescent microscope. 2.10. Cytochrome c detection Cells were cultured in complete medium for 24 h in 100 mm dishes (1  106 cells), then treated in serum free medium for 2 h. Cytochrome c was detected by Western blot analysis in mitochondrial and cytoplasmic fractions. Cells were harvested by centrifugation at 2500 rpm for 10 min at 4 °C. Pellets were resuspended in 50 ll of sucrose buffer (250 mM sucrose; 10 mM Hepes; 10 mM KCl; 1.5 mM MgCl2; 1 mM EDTA; 1 mM EGTA) containing 20 lg/ml aprotinin, 20 lg/ml leupeptin, 1 mM PMSF and 0.05% digitonine. Cells were incubated for 20 min at 4 °C and then centrifuged at 13,000 rpm for 15 min at 4 °C. The supernatant containing cytosolic protein fraction was transferred to new tubes and the resulting mitochondrial pellet was resuspended in 50 ll of lysis buffer (1% Triton X-100; 1 mM EDTA; 1 mM EGTA; 10 mM Tris–HCl, pH 7.4) containing 20 lg/ml aprotinin, 20 lg/ml leupeptin, 1 mM PMSF and then centrifuged at 13,000 rpm for 10 min at 4 °C. Equal amounts of proteins (10 lg) were resolved by 11% SDS/polyacrylamide gel as indicated in the Western blot analysis paragraph. 2.11. Detection of caspase 3 activity The activity of caspase 3 protease was measured using colorimetric activity assay kit (Sigma–Aldrich). Briefly, GC-2 cells were

52

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

cultured in complete medium for 24 h in 100 mm dishes (1  106 cells), then treated in serum free medium with E2, G1 and PPT (1 lM) for 6 h. Cells were harvested by centrifugation at 2000 rpm for 5 min at 4 °C, homogenized in lysis buffer and incubated on ice for 15 min. Cell debris were removed by centrifugation at 13,000 rpm for 15 min at 4 °C. Five microliters of cell lysates were combined with 2 mmol/l substrate (Ac-DEVD-pNA for caspase 3) and the volume brought up to 100 ll with the reaction buffer. Samples were covered and incubated at 37 °C overnight. Cleavage of Ac-DEVD-pNA by active caspase 3 resulted in the release of p-nitroanilide (pNA) into solution which was quantitated spectrophotometrically by measuring absorbance at 405 nm.

proliferation. Cells were treated for 48 h with E2, the specific ESR1 agonist (PPT) and the specific GPER agonist (G1). All ligands caused a dose-dependent reduction in GC-2 cell growth (Fig. 2A). To explain if this event was associated with a cell cycle arrest we investigated expression of G1 phase markers, such as cyclin D1 and cyclindependent kinase inhibitors WAF1/p21 (p21) and G2/M cyclins such as cyclin B1. Treatments with E2, G1 and PPT resulted in a decrease in cyclin D1 expression levels (Fig. 2B) and in a marked increase in p21 expression (Fig. 2C) compared with control cells. Moreover, the same treatments decreased cyclin B1 expression (Fig. 2D), as previously observed in primary PS (Chimento et al., 2010). These data are consistent with a G1 phase arrest, which was confirmed by flow cytometry analysis (data not shown).

2.12. RNA interference 3.3. E2, G1 and PPT induce DNA damage and apoptosis ESR1 siRNA, GPER siRNA and non targeting siRNA were purchased from Ambion. Cells were plated into 60 mm dishes at 1  106 cells for protein extraction, and into 24-well plates at 2  105 cells/well for proliferation assay and used for transfection 24 h later. siRNAs were transfected to a final concentration of 50 nM using Lipofectamine 2000 according to the manufacturer’s recommendations (Invitrogen). Proliferation was evaluated 48 h later. ESR1 and GPER-specific knockdown were checked by Western blot analysis of proteins extracted from cells transfected for 48 h. 2.13. Statistical analysis All experiments were conducted at least three times and the results were from representative experiments. Data were expressed as mean values + standard deviation (SD), and the statistical significance between control (basal) and treated samples was analyzed with SPSS10.0 statistical software. The unpaired Student’s t-test was used to compare two groups. P < 0.05 was considered statistically significant. 3. Results 3.1. Expression of estrogen receptors in GC-2 mouse spermatocytederived cell line The first aim of this study was to investigate ESR1, ESR2 and GPER expression in GC-2 cells. Both mRNA and protein levels of estrogen receptors were investigated. RT-PCR analysis demonstrated that GC-2 cells express both ERs isoforms as well as GPER (Fig. 1A and B). Protein expression analysis, using specific antibodies against the two receptor isoforms, confirmed mRNA data (Fig. 1C and D). TM3 immortalized mouse Leydig cell line was used as positive control for both ERs isoforms (Sirianni et al., 2007) while GC-1 mouse spermatogonial cell line was used as positive control for GPER (Sirianni et al., 2008). In addition, by immunofluorescence analysis using a specific GPER antibody, we defined the subcellular localization of GPER in GC-2 cells (Fig. 1E). The staining revealed a strong GPER immunoreactivity in the cytoplasm and plasma membrane (Fig. 1E, panel 5), whereas negative control cells showed immunonegative reaction for GPER (Fig. 1E, panel 2). 3.2. Activation of ESR1 and GPER inhibits GC-2 cell growth Recently, we have reported that E2 in rat pachytene spermatocytes, working through ESR1 and/or GPER, activates the rapid EGFR/ERK/c-Jun pathway, modulating the expression of genes involved in the balance between cellular proliferation and apoptosis (Chimento et al., 2010). Starting from these data, we decided to examine the effects of ESR1 and GPER activation on GC-2 cell

We then wanted to verify if the decrease in cell growth was associated with apoptosis. We first performed Annexin V assay, which is based on the high affinity of this protein for phosphatidylserine, a lipid absent from the outer membrane which translocates to the cell surface from the cytoplasmic side of the cell membrane during apoptosis. GC-2 cells treated with E2, G1 and PPT were strongly stained with Annexin V FITC (Fig. 3A). DAPI staining demonstrated that untreated GC-2 cells had round nuclei with regular contours and large in size. After treatment with E2, G1 and PPT cells showed nuclei shrunken and irregularly shaped or degraded with condensed DNA (Fig. 3B). Chromatin condensation was associated with DNA fragmentation (Fig. 3C). Gel electrophoresis of DNA extracted from GC-2 cells after 24 h treatment demonstrated the classic laddering pattern of inter-nucleosomal DNA fragmentation that was absent in control cells (Fig. 3C). To further confirm DNA damage after ESR1 and GPER activation Comet assay was used. Comet images of control GC-2 cells showed round cells without comet tail, that instead increased in length and amount of DNA after treatments with E2, G1 and PPT (Fig. 3D). 3.4. E2, G1 and PPT activate a mitochondria-dependent apoptotic pathway Since bcl-2 family members play pivotal roles in regulating the mitochondrial apoptotic pathway, bax and bcl-2 protein levels were detected by Western blot analysis. As seen in Fig. 4, the presence of E2, G1 and PPT increased bax (Fig. 4A), while decreased bcl-2 expression (Fig. 4B). Cytosolic translocation of cytochrome c has been proposed to be an essential component in the mitochondria-dependent pathway for apoptosis. Therefore, we examined if cytochrome c was released into the cytosol after ESR1 and GPER activation. Cell lysates were fractionated into cytosolic and mitochondrial fractions and analyzed by Western blot analysis (Fig. 4C and D). Cytochrome c levels in treated samples increased in the cytosolic fraction (Fig. 4C). While decreasing in the mitochondrial one (Fig. 4D). In order to confirm the involvement of ESR1 and GPER activation in the estrogen induced mitochondrialdependent apoptotic pathway, we evaluated cytochrome c expression into cytosolic and mitochondrial fractions after silencing the two receptors. The absence of the two proteins completely abolished E2 effect on cytochrome c release (Fig. 4E) and interfered with E2-dependent decrease in mitochondrial cytochrome c levels (Fig. 4F). When ESR1 or GPER were individually silenced E2 effect on cytochrome c release was only partially blocked, however, when both receptors were simultaneously silenced E2 completely lost its inhibitory effect (Fig. 4F). Since cytochrome c release from mitochondria into the cytosol triggers caspase activation (Hengartner, 2000), we then examined activation of the initiator caspase 9 and the executioner caspase 3 by Western blot analysis. After 6 h treatment with E2, G1 and PPT caspase 9 and caspase 3 (Fig. 4G)

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

53

Fig. 1. Expression of estrogen receptors in GC-2 cells. (A) ESR1 and ESR2 mRNA expression in GC-2 cells was analyzed by RT-PCR; TM3 cells were used as positive control; negative control (neg) contained water instead of cDNA. GAPDH was used as housekeeping gene. Size in base pair of amplified fragments is indicated. (B) GPER mRNA expression in GC-2 cells was analyzed by RT-PCR, GC-1 cells were used as positive control; negative control (neg) contained water instead of cDNA. GAPDH was used as housekeeping gene. Size in base pair of amplified fragments is indicated. (C) Western blot analysis of ESR1 and ESR2 was performed on 50 lg of total proteins extracted from GC-2 and TM3 cells. Blots are representative of three independent experiments with similar results. GAPDH was used as a loading control. (D) Western blot analysis of GPER was performed on 50 lg of total proteins extracted from GC-2 and GC-1 cells. GAPDH was used as loading control. Blots are representative of three independent experiments with similar results. (E) GC-2 cells were fixed and incubated with a polyclonal anti-GPER antibody (1:50) followed by incubation with goat anti-rabbit FITC-conjugated secondary antibody as described in Section 2. Cells were observed under a fluorescent microscope (magnification 400). Strong immunofluorescence was observed in the cytoplasm and membrane of GC-2 cells (panel 5). Negative control slides (neg) showed immunonegative reaction for GPER (panel 2). Propidium Iodide (PI) is localized in the nuclei of GC-2 cells (panels 3 and 6). Bright field (BF) contrast is shown for GC-2 cells (panels 1 and 4).

cleaved forms were visible. We evaluated also the effects of the three ligands on caspase 3 enzymatic activity, that was increased by 1.5-fold by E2, 1.7-fold by G1 and 2.5-fold by PPT (Fig. 4H). Once activated caspase 3 leads to the activation of poly (ADP-ribose) polymerase 1 (PARP-1), involved in the regulation of DNA repair (Dawson and Dawson, 2004). After 24 h treatment with E2, G1 and PPT we found that PARP-1 was activated, as seen by the presence of cleaved forms, (Fig. 4I). Importantly, when GC-2 cells were silenced with an ESR1 and/or GPER siRNA, the E2-induced PARP-1 activation was lost (Fig. 4J).

3.5. E2, G1 and PPT induce sustained MAPKs activation MAPK families, consisting of ERK, SAPK/JNK1/2 and p38, play central roles in cell proliferation, differentiation, survival, and apoptosis (Zhang and Liu, 2002). It has been originally shown that ERKs are important for cell survival, whereas JNKs and p38-MAPKs were deemed stress responsive and thus involved in apoptosis (Wada and Penninger, 2004). However, persistent or sustained ERK1/2 activation (Wong and Yan Cheng, 2005) and nuclear accumulation (Gonzalez et al., 1993; Zhang and Liu, 2002) are also involved in cell differentiation and death. Thus, we examined the activation of MAPK in response to E2, G1 and PPT. As shown in Fig. 5A, 10 min treatment with all three ligands determined ERK1/2, c-Jun, a direct JNK substrate, and p38 activation as seen

by increased phosphorylation, with maximum induction produced by 1 lM of each ligand. A time course study demonstrated a sustained activation of MAPK families, that was maintained higher than basal condition for up to 4 h (Fig. 5B). At this time point higher pERK1/2 levels were found in the nuclei of GC-2 cells as demonstrated by immunofluorescence staining (Fig. 5C). 3.6. ESR1 and GPER are both required for E2 effects on GC-2 cell proliferation In order to confirm that both ESR1 and GPER were required to produce E2-dependent effects on the inhibition of GC-2 cell proliferation, expression of the two receptors was silenced. Silencing ESR1 or GPER expression was able to partially block E2-dependent decrease in cell proliferation, when both receptors were simultaneously silenced E2 completely lost its inhibitory effect (Fig. 6A). Silencing of the two genes was assessed by Western blot analysis (Fig. 6B). 4. Discussion Spermatocytes represent a very interesting type of spermatogenic cells since can either proliferate or die through apoptosis (Sinha Hikim and Swerdloff, 1999) and represent a good target to control spermatogenesis. We have demonstrated that the

54

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

Fig. 2. Effects of ESR1 and GPER activators on GC-2 cell proliferation. (A) Cells were maintained for 24 h in complete medium and then treated for 48 h with increasing doses of E2, G1 and PPT (0.1, 1 and 10 lM). Proliferation was evaluated by [3H] thymidine incorporation analysis. Results were expressed as mean + SD of three independent experiments each performed in triplicate. (⁄, P < 0.05 and ⁄⁄, P < 0.01 compared with basal). (B–D) GC-2 cells were treated with E2, G1 and PPT (1 lM) for 48 h. Western blot analyses of cyclin D1 (B), p21 (C) and cyclin B1 (D) were performed on 50 lg of total proteins. Blots are representative of three independent experiments with similar results. GAPDH was used as a loading control. Graphs represent means of cyclin D1 (B), p21 (C) and cyclin B1 (D) optical densities normalized to GAPDH content of the same sample. Value of normalized optical density for basal sample was assumed as 1. (⁄, P < 0.01 compared with basal).

apoptotic process in these cells is induced by estradiol and requires activation of EGFR/ERK/c-Jun pathway and bax expression (Chimento et al., 2010). The study was performed using primary cultures of spermatocytes, that however, as major limit, could be maintained in cultures only for short times. Availability of a cell model that can reproduce the same behavior of primary cells would be helpful to further characterize apoptotic mechanisms activated by E2, indicating new targets to control cell proliferation. In this study using GC-2 cells we demonstrated that E2, through ESR1 and GPER, induces activation of signaling pathways involving MAPK families triggering cell apoptosis. We first evaluated ESR1, ESR2 and GPER expression in GC-2 cells and demonstrated an expression pattern similar to primary PS (Chimento et al., 2010). In addition using GC-2 cells we were able to show that the use of specific agonists for ESR1 and GPER caused a dose-dependent reduction in cell proliferation. It is well established that cell cycle progression is dynamically and strictly regulated by complexes containing cyclins and cyclin dependent kinases (CDKs), all of which are critical for the normal progression of cell cycle (John et al., 2001). G1/S phase progression is allowed by cyclin D1 and E (Sherr, 1995). Activity of these cyclins can be inhibited by the association with small inhibitory proteins,

the Cyclin-dependent Kinase Inhibitors (CKIs) such as p21 (Wade Harper et al., 1993). We found that after GPER and ESR1 activation cyclin D1 expression was reduced while p21 increased, confirming that cells do not bypass G1/S cell cycle check point as further demonstrated by flow cytometry analysis (data not shown). A similar effect was found in breast cancer cells, where GPER activation caused cell cycle arrest in G1 (Ariazi et al., 2010). In addition, we observed that the same treatment decreased cyclin B1 expression in according with previous data obtained in PS primary cultures (Chimento et al., 2010). It exists a strict relationship between G1 cyclins and cell apoptosis, in fact G1 cyclins activity is essential for G1 exit and apoptosis escape (Chiarugi et al., 1994). Apoptosis has long been recognized as a significant feature of mammalian spermatogenesis (Sinha Hikim and Swerdloff, 1999). Here we demonstrated that the effect of E2, G1 and PPT on GC-2 cell cycle was associated with initiation of an apoptotic mechanism. An early indicator of apoptosis is the rapid translocation and accumulation of the membrane phospholipid phosphatidylserine (PhSer) from cytoplasmic interface to the extracellular surface (Chan et al., 1998). This loss of membrane asymmetry can be detected by utilizing the binding properties of Annexin V for negatively charged phospholipids

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

55

Fig. 3. Effects of ESR1 and GPER activation on GC-2 apoptosis and nuclei morphology. (A) GC-2 cells were left untreated (bs) or treated with E2, G1 and PPT (1 lM) for 12 h. Annexin FITC staining assay was performed as described in Section 2. Cells were observed under a fluorescent microscope (magnification 400). Images are from a representative experiment. (B–D) GC-2 cells were left untreated (basal) or treated with E2, G1 and PPT (1 lM) for 24 h. (B) After treatment GC-2 cells were fixed with paraformaldehyde, dyed with DAPI and observed under fluorescent microscope (magnification 400). Arrows indicate condensed nuclei. Images are from a representative experiment. (C) After treatment DNA was extracted from cells and analyzed on a 1.5% agarose gel. (D) After treatment DNA damage was analyzed by Comet assay. Cells were observed under a fluorescent microscope (magnification 1000). Images are from a representative experiment.

including PhSer in early apoptotic cells (Koopman et al., 1994). Here, we have shown that E2, G1 and PPT strongly induce positivity for Annexin V FITC staining. Moreover we demonstrated the induction of apoptosis through DAPI staining, that evidenced nuclei morphological changes and through comet and DNA fragmentation assay that evidenced laddering pattern of inter-nucleosomal DNA. It is generally accepted the theory that apoptosis can be induced by that extrinsic (Kim et al., 2006) and intrinsic (Fadeel and Orrenius, 2005) mechanisms. In the latter, bcl-2 family of proteins plays a central role (Cory and Adams, 2002). This family consists of both pro-(bax, bad, bak, bid) and anti-apoptotic (bcl-2, bcl-xl) proteins that modulate the execution phase of the

cell death pathway. Expression of these proteins changes in testicular germ cell apoptosis (Yan et al., 2000). Bax exerts pro-apoptotic activity by cytochrome c translocation from the mitochondria to cytosol (Antonsson et al., 2000) where it binds to apoptotic protease-activating factor-1 (Apaf-1) (Wang, 2001), which in turn binds to procaspase 9 via the caspase recruitment domain at the amino terminus in the presence of deoxy-ATP, resulting in activation of the initiator caspase 9 (Kuida et al., 1998) and subsequent proteolytic activation of executioner caspase 3 (Wilson, 1998). The active caspase 3 is then involved in the cleavage of a set of proteins including poly-(ADP) ribose polymerase (PARP) (Soldani and Scovassi, 2002). Bcl-2, instead, exerts its anti-apoptotic activity,

56

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

Fig. 4. Activation of mitochondrial apoptotic pathway by E2, G1 and PPT. (A and B) Proteins extracted from GC-2 cells treated with E2, G1 and PPT (1 lM) for 24 h were subjected to Western blot analysis for bax and bcl-2. GAPDH was used as a loading control. (C and D) GC-2 cells were left untreated (basal) or treated with E2, G1 and PPT (1 lM) for 2 h. Cytochrome c (Cyt c) levels in cytosolic (C) and mitochondrial (D) fractions were detected by Western blot analysis. Protein expression in each lane was normalized to the GAPDH content. Graphs represent means of Cyt c optical densities normalized to GAPDH content of the same sample. Value of normalized optical density for basal sample was assumed as 1. (⁄, P < 0.01 compared with basal). (E and F) GC-2 cells were transfected with ESR1, GPER, non targeting (control siRNA) siRNA (50 nM) as indicated. Seventy hours after transfection cells were treated for an additional 2 h with E2 (1 lM). Cytochrome c (Cyt c) levels in cytosolic (C) and mitochondrial (D) fractions were detected by Western blot analysis. Protein expression in each lane was normalized to the GAPDH content. Graphs represent means of Cyt c optical densities normalized to GAPDH content of the same sample. Value of normalized optical density for basal sample was assumed as 1. (G and H) GC-2 cells were treated with E2, G1 and PPT (1 lM) for 6 h. (G) Appearance of caspase 9 and caspase 3 cleaved form was determined by Western blot analysis. GAPDH was used as a loading control. (H) Caspase activity was analyzed by caspase 3 colorimetric assay kit as indicated in the Section 2. Data were expressed as mean + SD of three independent experiments. (⁄, P < 0.05 compared with basal). (I) GC-2 cells were treated with E2, G1, PPT (1 lM) for 24 h; 50 lg of total proteins were analyzed by Western blot for PARP-1 activation. GAPDH was used as a loading control. (J) GC-2 cells were transfected with ESR1, GPER, non targeting (control siRNA) siRNA (50 nM) as indicated. Forty-eight hours after transfection cells were treated for an additional 24 h with E2 (1 lM). Fifty lg of total proteins were analyzed by Western blot for PARP activation. GAPDH was used as a loading control.

at least in part, by inhibiting the translocation of bax to the mitochondria (Wang, 2001). Evidence for a crucial role of bax and bcl-2 in spermatogenesis comes from transgenic mice over-expressing bcl-2 or bax-deficient male mice which are infertile, an event attributed to the disruption of spermatogenesis (Rodriguez et al., 1997). GC-2 cells responded to E2 by increasing bax and concom-

itantly decreasing bcl-2 expression, increasing cytochrome c release into the cytosol and activating caspase 9, caspase 3 and PARP, demonstrating the ability of estradiol to activate an intrinsic apoptotic mechanism. Silencing of both ESR1 and GPER genes confirmed the role of the two receptors in mitochondria-dependent apoptotic pathway. In fact, the reduced expression of both

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

57

Fig. 5. Effects of E2, G1 and PPT on ERK1/2, c-Jun and p38 activation. (A) GC-2 cells were treated for 10 min with the indicated concentrations of E2, G1 and PPT. Western blot analyses of pERK1/2, p-cJun and phospho p38 were performed on 50 lg of total proteins. ERK1/2, cJun or p38 were used as a loading control. Blots are representative of three independent experiments with similar results. (B) GC-2 were treated for the indicated times with E2, G1 and PPT (1 lM). Western blot analyses of pERK1/2, p-cJun and phospho-p38 were performed on 50 lg of total proteins. ERK1/2, cJun or p38 were used as a loading control. Blots are representative of three independent experiments with similar results. (C) GC-2 cells were treated with E2, G1 and PPT (1 lM) for 4 h. ERK activation was determined by green immunofluorescence as indicated in Section 2. Nuclei were stained with DAPI (blue). Cells were observed under a fluorescent microscope (magnification 200).

receptors completely abolished effects of E2 on cytochrome c release in the cytosol and on PARP-1 activation. Initiation of apoptosis occurs after activation of different signaling pathways. Mitogen-activated protein kinase (MAPK) cascades have been shown to play a key role in converting extracellular signals into cellular responses, such as cell proliferation, cell differentiation, cell motility and cell death (Pearson et al., 2001). In mammalian cells, three MAPK families have been clearly characterized: namely classical MAPK (also known as ERK), c-Jun N-terminal kinase/stress-activated protein kinase (JNK/SAPK) and p38 kinase (Zhang and Liu, 2002). The signaling cascades involving JNK and p38, activated by extracellular stress signals, are involved in cell differentiation and apoptosis (Xia et al., 1995). Importantly, a previous study utilizing immunohistochemistry demonstrated that phospho-p38 localized mainly in PS and some Sertoli cells in control testes from 21-day-old rats and its activation was further

increased following heat-induced apoptosis (Lizama et al., 2009). Previous studies have demonstrated that transient activation of ERK1/2 plays a pivotal role in cell proliferation and that sustained ERK1/2 activation induces cell cycle arrest (Adachi et al., 2002) and death (Chen et al., 2005). In the present study we showed that E2, G1 and PPT caused a rapid, however sustained (6 h), ERK1/2 activation, as demonstrated by the presence of their phosphorylated form in GC-2 cell nuclei. Similarly, both p38 and c-Jun, a direct target of JNK, phosphorylation was rapid and sustained following GPER and ESR1 activation. The involvement of both receptors in apoptosis was confirmed by our last set of experiments demonstrating that silencing of ESR1 and GPER gene expression caused the loss of E2 inhibitory effect on cell proliferation. It remains to establish the role of ESR2 in PS, that we are currently investigating. Preliminary results indicate that DPN (selective agonist of ESR2) activates ERK1/2 but causes an increase in cyclin B1 and a decrease

58

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59

Fig. 6. Effects of ESR1 and GPER silencing on E2-dependent inhibition of cell proliferation. (A) GC-2 cells were transfected with ESR1, GPER, non targeting (control siRNA) siRNA as indicated. Twenty-four hours after transfection cells were treated for an additional 48 h with E2 (1 lM). Proliferation was evaluated by [3H]thymidine incorporation assay. Results were expressed as mean + SD of three independent experiments each performed in triplicate. (B) GC-2 cells were transfected with ESR1, GPER, non targeting (control siRNA) siRNA as indicated. Seventy-two hours after transfection cells were lysed and subject to Western blot analyses for ESR1 and GPER. GAPDH was used as a loading control. Results are representative of three independent experiments.

in Bax mRNA expression suggesting that in PS ESR2 plays a role similar to the one played in round spermatids (Chimento et al., 2011). This phenomenon can be explained by the ability of ESR2 in activating different E2-dependent pathways as it was shown in other cellular system (Matthews and Gustafsson, 2003). Stimulation of both pathways by E2 determines a prevalence of apoptotic signals. This could be explained by the higher ESR1 and GPER expression compared to ESR2. Taken together, the present study indicates that GC-2 cells express ESR1 and GPER, which are both involved in E2-dependent apoptotis as previously demonstrated in primary PS. Moreover the use of GC-2 cells allowed to further characterize the E2-dependent molecular pathway associated with apoptosis of PS. This involves activation of MAPK families ERK1/2, JNK and p38, followed by activation of intrinsic apoptotic pathway determining bax up-regulation, cytochrome c release, caspase 9 and 3 activation and DNA fragmentation. The definition of the molecular components of germ cells apoptosis will help us to better understand the consequences of the exposure to environmental estrogens, to provide new potential targets for the development of a non-androgen male contraceptive or for the development of novel therapeutic regimens to control germ cell tumors. Disclosure statement The authors have nothing to disclose. Acknowledgements This work was supported by Associazione Italiana per la Ricerca sul Cancro (AIRC) project no. IG10344.

References Adachi, T., Kar, S., Wang, M., Carr, B.I., 2002. Transient and sustained ERK phosphorylation and nuclear translocation in growth control. J. Cell. Physiol. 192, 151–159. Antonsson, B., Montessuit, S., Lauper, S., Eskes, R., Martinou, J.C., 2000. Bax oligomerization is required for channel-forming activity in liposomes and to trigger cytochrome c release from mitochondria. Biochem. J. 345, 271–278. Ariazi, E.A., Brailoiu, E., Yerrum, S., Shupp, H.A., Slifker, M.J., Cunliffe, H.E., Black, M.A., Donato, A.L., Arterburn, J.B., Oprea, T.I., Prossnitz, E.R., Dun, N.J., Jordan, V.C., 2010. The G Protein-Coupled Receptor GPR30 inhibits proliferation of estrogen receptor-positive breast cancer cells. Cancer Res. 70, 1184–1194. Billig, H., Furuta, I., Rivier, C., Tapanainen, J., Parvinen, M., Hsueh, A.J., 1995. Apoptosis in testis germ cells: developmental changes in gonadotropin dependence and localization to selective tubule stages. Endocrinology 136, 5– 12. Bois, C., Delalande, C., Nurmio, M., Parvinen, M., Zanatta, L., Toppari, J., Carreau, S., 2010. Age- and cell-related gene expression of aromatase and estrogen receptors in the rat testis. J. Mol. Endocrinol. 45, 147–159. Brailoiu, E., Dun, S.L., Brailoiu, G.C., Mizuo, K., Sklar, L.A., Oprea, T.I., Prossnitz, E.R., Dun, N.J., 2007. Distribution and characterization of estrogen receptor G protein-coupled receptor 30 in the rat central nervous system. J. Endocrinol. 193, 311–321. Carreau, S., Delalande, C., Galeraud-Denis, I., 2009. Mammalian sperm quality and aromatase expression. Microsc. Res. Tech. 72, 552. Carreau, S., Lambard, S., Delalande, C., is-Galeraud, I., Bilinska, B., Bourguiba, S., 2003. Aromatase expression and role of estrogens in male gonad: a review. Reprod. Biol. Endocrinol. 1, 35. Chagin, A.S., Savendahl, L., 2007. GPR30 estrogen receptor expression in the growth plate declines as puberty progresses. J. Clin. Endocrinol. Metab. 92, 4873–4877. Chan, A., Reiter, R., Wiese, S., Fertig, G., Gold, R., 1998. Plasma membrane phospholipid asymmetry precedes DNA fragmentation in different apoptotic cell models. Histochem. Cell Biol. 110, 553–558. Chen, J.-R., Plotkin, L.I., Aguirre, J.I., Han, L., Jilka, R.L., Kousteni, S., Bellido, T., Manolagas, S.C., 2005. Transient versus sustained phosphorylation and nuclear accumulation of ERKs underlie anti-versus pro-apoptotic effects of estrogens. J. Biol. Chem. 280, 4632–4638. Chiarugi, V., Magnelli, L., Cinelli, M., Basi, G., 1994. Apoptosis and the cell cycle. Cell. Mol. Biol. Res. 40, 603–612. Chimento, A., Sirianni, R., Delalande, C., Silandre, D., Bois, C., Ando’, S., Maggiolini, M., Carreau, S., Pezzi, V., 2010. 17b-estradiol activates rapid signaling pathways

A. Chimento et al. / Molecular and Cellular Endocrinology 355 (2012) 49–59 involved in rat pachytene spermatocytes apoptosis through GPR30 and ERa. Mol. Cell Endocrinol. 320, 136–144. Chimento, A., Sirianni, R., Zolea, F., Bois, C., Delalande, C., Ando’, S., Maggiolini, M., Aquila, S., Carreau, S., Pezzi, V., 2011. Gper and ESRs are expressed in rat round spermatids and mediate oestrogen-dependent rapid pathways modulating expression of cyclin B1 and Bax. Int. J. Androl. 34, 420–429. Cory, S., Adams, J.M., 2002. The Bcl2 family: regulators of the cellular life-or-death switch. Nat. Rev. Cancer. 2, 647–656. D’Souza, R., Gill-Sharma, M.K., Pathak, S., Kedia, N., Kumar, R., Balasinor, N., 2005. Effect of high intratesticular estrogen on the seminiferous epithelium in adult male rats. Mol. Cell Endocrinol. 241, 41–48. Dawson, V.L., Dawson, T.M., 2004. Deadly conversations: nuclear-mitochondrial cross-talk. J. Bioenerg. Biomembr. 36, 287–294. Delbes, G., Levacher, C., Pairault, C., Racine, C., Duquenne, C., Krust, A., Habert, R., 2004. Estrogen receptor {beta}-mediated inhibition of male germ cell line development in mice by endogenous estrogens during perinatal life. Endocrinology 145, 3395–3403. Ebling, F.J.P., Brooks, A.N., Cronin, A.S., Ford, H., Kerr, J.B., 2000. Estrogenic induction of spermatogenesis in the hypogonadal mouse. Endocrinology 141, 2861–2869. Fadeel, B., Orrenius, S., 2005. Apoptosis: a basic biological phenomenon with wideranging implications in human disease. J. Int. Med. 258, 479–517. Funakoshi, T., Yanai, A., Shinoda, K., Kawano, M.M., Mizukami, Y., 2006. G proteincoupled receptor 30 is an estrogen receptor in the plasma membrane. Biochem. Biophys. Res. Comm. 346, 904–910. Gancarczyk, M., Paziewska-Hejmej, A., Carreau, S., Tabarowski, Z., Bilin´ska, B., 2004. Dose- and photoperiod-dependent effects of 17beta-estradiol and the antiestrogen ICI 182,780 on testicular structure, acceleration of spermatogenesis, and aromatase immunoexpression in immature bank voles. Acta Histochem. 106, 269–278. Gonzalez, F., Seth, A., Raden, D., Bowman, D., Fay, F., Davis, R., 1993. Serum-induced translocation of mitogen-activated protein kinase to the cell surface ruffling membrane and the nucleus. J. Cell Biol. 122, 1089–1101. Hengartner, M.O., 2000. The biochemistry of apoptosis. Nature 407, 770–776. Hofmann, M.C., Hess, R.A., Goldberg, E., Millán, J.L., 1994. Immortalized germ cells undergo meiosis in vitro. Proc. Natl. Acad. Sci. USA 91, 5533–5537. Isensee, J., Meoli, L., Zazzu, V., Nabzdyk, C., Witt, H., Soewarto, D., Effertz, K., Fuchs, H., Gailus-Durner, V., Busch, D., Adler, T., de angelis, M.H., Irgang, M., Otto, C., Noppinger, P.R., 2009. Expression pattern of G Protein-Coupled Receptor 30 in lacz reporter mice. Endocrinology 150, 1722–1730. John, P., Mews, M., Moore, R., 2001. Cyclin/Cdk complexes: their involvement in cell cycle progression and mitotic division. Protoplasma 216, 119–142. Kim, R., Emi, M., Tanabe, K., Murakami, S., Uchida, Y., Arihiro, K., 2006. Regulation and interplay of apoptotic and non-apoptotic cell death. J. Pathol. 208, 319–326. Koopman, G., Reutelingsperger, C., Kuijten, G., Keehnen, R., Pals, S., van Oers, M., 1994. Annexin V for flow cytometric detection of phosphatidylserine expression on B cells undergoing apoptosis. Blood 84, 1415–1420. Kuida, K., Haydar, T.F., Kuan, C.-Y., Gu, Y., Taya, C., Karasuyama, H., Su, M.S.S., Rakic, P., Flavell, R.A., 1998. Reduced apoptosis and cytochrome c mediated caspase activation in mice lacking caspase 9. Cell 94, 325–337. Lee, M., Kwon, J., Chung, M.K., 2003. Enhanced prediction of potential rodent carcinogenicity by utilizing comet assay and apoptotic assay in combination. Mutat. Res.-Gen. Tox. En. 541, 9–19. Levin, E.R., 2008. Rapid signaling by steroid receptors. Am. J. Physiol.- Reg. I 295, R1425–R1430. Lizama, C., Lagos, C.F., Lagos-Cabré, R., Cantuarias, L., Rivera, F., Huenchuñir, P., Pérez-Acle, T., Carrión, F., Moreno, R.D., 2009. Calpain inhibitors prevent p38 MAPK activation and germ cell apoptosis after heat stress in pubertal rat testes. J. Cell. Physiol. 221, 296–305. Lucas, T.s.F.G., Royer, C., Siu, E.R., Lazari, M.F., Porto, C.S., 2010. Expression and signaling of G Protein-Coupled Estrogen Receptor 1 (GPER) in rat Sertoli cells. Biol. Reprod. 83, 307–317. Matthews, J., Gustafsson, J.A., 2003. Estrogen signaling: a subtle balance between ER alpha and ER beta. Mol. Interv. 3, 281–292.

59

Mishra, D.P., Shaha, C., 2005. Estrogen-induced spermatogenic cell apoptosis occurs via the mitochondrial pathway: role of superoxide and nitric oxide. J. Biol. Chem. 280, 6181–6196. Pearson, G., Robinson, F., Beers Gibson, T., Xu, B.-e., Karandikar, M., Berman, K., Cobb, M.H., 2001. Mitogen-Activated Protein (MAP) kinase pathways: regulation and physiological functions. Endocr. Rev. 22, 153–183. Pentikäinen, V., Erkkilä, K., Suomalainen, L., Parvinen, M., Dunkel, L., 2000. Estradiol acts as a germ cell survival factor in the human testis in vitro. J. Clin. Endocrinol. Metab. 85, 2057–2067. Perrard, M.H., Durand, P., 2009. Redundancy of the effect of TGFbeta1 and beta-NGF on the second meiotic division of rat spermatocytes. Microsc. Res. Tech. 72, 596–602. Revankar, C.M., Cimino, D.F., Sklar, L.A., Arterburn, J.B., Prossnitz, E.R., 2005. A transmembrane intracellular estrogen receptor mediates rapid cell signaling. Science 307, 1625–1630. Rodriguez, I., Ody, C., Araki, K., Garcia, I., Vassalli, P., 1997. An early and massive wave of germinal cell apoptosis is required for the development of functional spermatogenesis. EMBO J. 16, 2262–2270. Sakamoto, H., Matsuda, K.i., Hosokawa, K., Nishi, M., Morris, J.F., Prossnitz, E.R., Kawata, M., 2007. Expression of GPR30, a G protein-coupled membrane estrogen receptor, in oxytocin neurons of the rat paraventricular and supraoptic nuclei. Endocrinology 148, 5842–5850. Sherr, C., 1995. Mammalian G1 cyclins and cell cycle progression. P. Assoc. Am. Physician. 107, 181–186. Sinha Hikim, A.P., Swerdloff, R.S., 1999. Hormonal and genetic control of germ cell apoptosis in the testis. Rev. Reprod. 4, 38–47. Sirianni, R., Chimento, A., Malivindi, R., Mazzitelli, I., Andò, S., Pezzi, V., 2007. Insulin-like growth factor-I, regulating aromatase expression through steroidogenic factor 1, supports estrogen-dependent tumor Leydig cell proliferation. Cancer Res. 67, 8368–8377. Sirianni, R., Chimento, A., Ruggiero, C., De Luca, A., Lappano, R., Andò, S., Maggiolini, M., Pezzi, V., 2008. The novel estrogen receptor, G Protein-Coupled Receptor 30, mediates the proliferative effects induced by 17{beta}-estradiol on mouse spermatogonial GC-1 cell line. Endocrinology 149, 5043–5051. Soldani, C., Scovassi, A.I., 2002. Poly(ADP-ribose) polymerase-1 cleavage during apoptosis: an update. Apoptosis 7, 321–328. Tice, R.R., Agurell, E., Anderson, D., Burlinson, B., Hartmann, A., Kobayashi, H., Miyamae, Y., Rojas, E., Ryu, J.C., Sasaki, Y.F., 2000. Single cell gel/comet assay: guidelines for in vitro and in vivo genetic toxicology testing. Envirol. Mol. Mutagen. 35, 206–221. Wada, T., Penninger, J.M., 2004. Mitogen-activated protein kinases in apoptosis regulation. Oncogene 23, 2838–2849. Wade Harper, J., Adami, G.R., Wei, N., Keyomarsi, K., Elledge, S.J., 1993. The p21 Cdkinteracting protein Cip1 is a potent inhibitor of G1 cyclin-dependent kinases. Cell 75, 805–816. Wang, C., Prossnitz, E.R., Roy, S.K., 2007. Expression of G Protein-Coupled Receptor 30 in the hamster ovary: differential regulation by gonadotropins and steroid hormones. Endocrinology 148, 4853–4864. Wang, X., 2001. The expanding role of mitochondria in apoptosis. Gen. Dev. 15, 2922–2933. Wilson, M.R., 1998. Apoptosis: unmasking the executioner. Cell Death Differ. 5, 646–652. Wong, C.H., Yan Cheng, C., 2005. Mitogen-activated protein kinases, adherens junction dynamics, and spermatogenesis: a review of recent data. Dev. Biol. 286, 1–15. Xia, Z., Dickens, M., Raingeaud, J., Davis, R.J., Greenberg, M.E., 1995. Opposing effects of ERK and JNK-p38 MAP kinases on apoptosis. Science 270, 1326–1331. Yan, W., Suominen, J., Samson, M., Jégou, B., Toppari, J., 2000. Involvement of Bcl-2 family proteins in germ cell apoptosis during testicular development in the rat and pro-survival effect of stem cell factor on germ cells in vitro. Mol. Cell. Endocrinol. 165, 115–129. Zhang, W., Liu, H.T., 2002. MAPK signal pathways in the regulation of cell proliferation in mammalian cells. Cell Res. 12, 9–18.