Abstract
Spermatogenesis is dependent on the ability of Sertoli cells to form mature junctions that maintain a unique environment within the seminiferous epithelium. Adjacent Sertoli cells form a junctional complex that includes classical adherens junctions and testis-specific ectoplasmic specialisations (ES). The regulation of inter-Sertoli cell junctions by the two main endocrine regulators of spermatogenesis, FSH and testosterone, is unclear. This study aimed to investigate the effects of FSH and testosterone on inter-Sertoli cell adherens junctions (as determined by immunolocalisation of cadherin, catenin and actin) and ES junctions (as determined by immunolocalisation of espin, actin and vinculin) in cultured immature Sertoli cells and GnRH-immunised adult rat testes given FSH or testosterone replacement in vivo. When hormones were absent in vitro, adherens junctions formed as discrete puncta between interdigitating, finger-like projections of Sertoli cells, but ES junctions were not present. The adherens junction puncta included actin filaments that were oriented perpendicularly to the Sertoli cell plasma membrane, but were not associated with the intermediate filament protein vimentin. When FSH was added in vitro, ES junctions formed, and adjacent adherens junction puncta fused into extensive adherens junction belts. After hormone suppression in vivo, ES junctions were absent, while FSH replacement restored ES junctions, as confirmed by electron microscopy and confocal analysis of ES-associated proteins. Testosterone alone did not affect adherens junctions or ES in vitro or in vivo. We conclude that FSH can regulate the formation of ES junctions and stimulate the organisation and orientation of extensive adherens junctions in Sertoli cells.
Introduction
Spermatogenesis is defined as the process of male germ cell development, and involves the division, chromosome reduction and morphological differentiation of diploid stem-line germ cells to produce haploid spermatozoa (Steinberger & Steinberger 1975). Sertoli cells are the somatic cells of the testis, serving to nurture germ cells as they proceed through spermatogenesis (Steinberger & Steinberger 1975). Sertoli cells are polarised epithelial cells that interact with one another via a unique and large complex of basolaterally located junctions (Vogl et al. 1993). This complex consists of various junction types, including adherens junctions (AJ), desmosome-like junctions, tight junctions and gap junctions (Russell & Peterson 1985, Vogl et al. 1993), and is further characterised by the presence of testis-specific Sertoli cell structures termed ‘ectoplasmic specialisations’ (ES) (Vogl et al. 2000). The entire inter-Sertoli cell junctional complex regulates the movement of substances into and out of the seminiferous epithelium, thus forming the basis of the blood–testis barrier. The formation of this barrier is essential for normal germ cell development (Steinberger & Steinberger 1975, Weber et al. 1988).
Typical AJs are found between Sertoli cells and are akin to those found in all classical epithelia. AJs are characterised by the presence of transmembrane, calcium-dependent cadherins that link to the actin cytoskeleton via catenins (see Lui et al. (2003) for recent review) and are visible by electron microscopy as focal opposing subsurface densities on plasma membranes (Russell & Peterson 1985, Byers et al. 1993). In classical epithelia, AJs are formed initially as AJ puncta where actin filaments terminate perpendicularly to the plasma membrane, and, upon appropriate stimulation, reorganise to form AJ belts in which actin filaments are oriented parallel to the plasma membrane (Krendel & Bonder 1999, Vasioukhin & Fuchs 2001). Although the presence of inter-Sertoli cell AJs has been well documented (reviewed in Russell & Peterson 1985 and Lui et al. 2003), the regulation of AJ puncta and AJ belt formation in Sertoli cells has not yet been described.
The ES is generally regarded as a testis-specific, actin-based adhesion junction (Vogl et al. 2000, Toyama et al. 2003, Mruk & Cheng 2004); however, they exhibit molecular, ultrastructural and regulatory differences from classical AJs. ES junctions are found only in the testis, and are a Sertoli cell cytoskeletal structure characterised by hexagonally packed bundles of actin filaments, oriented parallel to the Sertoli cell plasma membrane, and sandwiched between the plasma membrane and cisternae of the endoplasmic reticulum (Russell 1977, Vogl et al. 1991). The packing of actin filaments into hexagonal bundles is achieved by the actin-bundling protein espin (Bartles et al. 1996), while vinculin, also localised to the layer of actin filaments (Pfeiffer & Vogl 1991), is thought to link the actin filaments of the ES to transmembrane molecules (Pokutta & Weis 2002). Furthermore, actin filaments at ES junctions are arranged in a regular and unipolar manner, in contrast to classical AJs (Vogl et al. 1993), and do not exhibit contractile properties (Vogl et al. 1993). ES junctions are present at two sites within the seminiferous epithelium; between adjacent Sertoli cells (basal ES junctions), and between Sertoli cells and elongating and elongated spermatids (apical ES junctions). Basal ES junctions are characterised by two opposing ES structures (Vogl et al. 2000).
The transmembrane and intercellular adhesion molecules that comprise the putative adhesive domain of ES junctions are unclear, although β1-integrin is a likely candidate (Mulholland et al. 2001). The nectin-afadin system may also mediate adhesion at sites of apical ES junctions (Ozaki-Kuroda et al. 2002). In contrast, cadherins, the classical transmembrane adhesion molecules found at AJs (Johnson & Boekelheide 2002), have been reported to be absent from ES junctions (Mulholland et al. 2001, Johnson & Boekelheide 2002), although a recent study suggests that N-cadherin is present at sites consistent with ES junctions (Lee et al. 2004).
Although it is well known that testosterone is essential for spermatogenesis, a consensus is developing that follicle-stimulating hormone (FSH) is also required for quantitatively normal spermatogenesis in the rodent (see McLachlan et al. (2002) and O’Donnell et al.(2005) for review). Transgenic male mice with targeted disruption of the FSHβ gene are fertile (Kumar et al. 1997), while male mice lacking a functional FSH receptor show some impairment of fertility (Dierich et al. 1998, Krishnamurthy et al. 2000). However, a closer examination of the phenotypes of these mice revealed defects in the ability of Sertoli cells to support germ cells in FSHβ-null mice (Wreford et al. 2001) and defects to germ cells, such as elongated spermatids (Krishnamurthy et al. 2000) and Sertoli cells (Grover et al. 2004), in FSH receptor-null mice. Further support for a role for FSH in spermatogenesis comes from a study demonstrating that short-term (1-week) passive immunoneutralisation of circulating FSH in normal rats decreased germ cell numbers and increased germ cell apoptosis in a setting of normal serum and testicular testosterone (Meachem et al. 1999).
In order to begin to understand the relationship between hormones, spermatogenesis and Sertoli cell junctions, earlier studies assessed the effects of FSH and testosterone on Sertoli cell junctional structures that contain actin and vinculin in adult rats (Muffly et al. 1993, 1994). The reduction of luteinizing hormone (LH) and FSH by hypophysectomy resulted in an abnormal pattern of actin and vinculin staining in the seminiferous epithelium (Muffly et al. 1993, 1994). The subsequent restoration of FSH, but not testosterone (Muffly et al. 1993, 1994), re-established the normal organisation of these junctional molecules, which correlated with the restoration of spermatogenesis. These data indicate that FSH promotes the ability of Sertoli cells to form normal junctions, both with themselves and with other germ cells, and this, in turn, is important for normal spermatogenesis (Muffly et al. 1993, 1994). FSH has also been shown to regulate the formation of integrin-containing junctions between developing Sertoli cells (Salanova et al. 1998). However, the regulation, mechanism of formation and interdependence of different junction types at the inter-Sertoli cell junctional complex remain largely unknown.
It is hypothesised that FSH and/or testosterone regulate the formation of mature adhesion junctions, including AJs and ES junctions, between Sertoli cells. Hence, the aim of this study was to examine the role of FSH and testosterone in the formation of classical cadherin/catenin-based AJs and the Sertoli cell-specific ES junction, containing actin and espin, in immature rat Sertoli cells in vitro, and in adult rats in vivo. The gonadotrophin-releasing hormone (GnRH)-immunised rodent model was chosen, as it is analogous to current androgen-based male contraceptive methods (McLachlan et al. 2002).
Materials and Methods
Animals
Male Sprague-Dawley rats (19–21 or 75–85 days old) obtained from Monash Animal Services (Monash University, Clayton, Australia) were maintained at 20 °C in a fixed 12-h light–dark cycle with free access to food and water. Animal experimentation was approved by the Monash Medical Centre animal ethics committee.
In vitro studies
Sertoli cell isolation and culture
Immature Sertoli cells for use in culture were isolated by previously published protocols (Cameron & Muffly 1991, Perryman et al. 1996). Briefly, twelve 19–21-day-old male Sprague Dawley rats were killed by CO2 asphyxiation. Testes were removed and decapsulated, and seminiferous tubules were finely chopped. Interstitial tissue was removed by digestion with trypsin (2.5 mg/ml; Sigma)/DNase (50 μg/ml; Sigma) dissolved in Dulbecco’s PBS (DPB; Gibco), using an orbital shaking air bath (25 min, 37 °C, 85 r.p.m.). Digestion was halted by the addition of 10% fetal calf serum (FCS) (Trace Biosciences, Sydney, Australia)/ DNase (10 μg/ml) in DPB. Seminiferous tubules were separated from interstitial fragments by unit gravity sedimentation, followed by a series of wash/sediment cycles in 10% FCS/DNase (10 μg/ml), and three washes in DNase (20 μg/ml) in DPB. Seminiferous tubules were then resuspended with 1% BSA (Sigma)/DNase (10 μg/ml) in DPB and processed in a glass tissue homogeniser to generate a single-cell suspension. Cells were filtered through 80 μm mesh, pelleted by centrifugation (100 g for 5 min), washed twice with DPB, and resuspended with 10 ml culture media (see below).
Isolated Sertoli cells were plated (1 × 106 cells/ml per well; 0.5 × 106 cells per cm2) into 24-well culture plates (Costar, Acton, MA, USA) containing glass cover-slips (13 mm diameter; Knittel Gläser, Braunschweig, Germany) previously coated with Matrigel (1.97 mg/ml in DMEM; 20 μl/well, excess removed; BD Biosciences Bedford, MA, USA). This plating concentration of Sertoli cells results in the seeding of Sertoli cells at a final density of approximately 20 Sertoli cells/10 000 μm2, as assessed by stereological counting of Sertoli cell nuclei after culture (data not shown).
After plating, cells were incubated for 72 h at 37 °C in a humidified 5% CO2/95% air incubator, and then hypotonically shocked with 10% culture media in water for 45 s to remove contaminating germ cells from the Sertoli cell monolayer, and replaced in culture medium. Sertoli cells were then allowed to recover for a period of 24 h at 37 °C, 5% CO2/95% air. Culture media were replaced, hormones added, and cells cultured for a further 48 h at 37 °C, 5% CO2/95% air, after which they were fixed and processed for immunocytochemistry as described below.
Culture media consisted of DMEM:F12 (mixed 1:1 v/v; Trace Biosciences, Sydney, Australia) supplemented with 1 × non-essential amino acids (Trace Biosciences), l-glutamine (1 mM, Trace Biosciences), sodium bicarbonate (0.12% w/v, Trace Biosciences) and penicillin/streptomycin/fungizone (200 U/ml, 200 μg/ ml, 500 ng/ml respectively; CSL Limited, Melbourne, Australia) with final pH 7.2–7.4. Prior to use, BSA (1% w/v; Sigma), HEPES buffer (10 mM) and penicillin/ streptomycin/fungizone (as above) were added to the culture media.
In vitro hormone treatments
Where indicated, human recombinant (hr)FSH (Puregon, N.V. Organon, Oss, The Netherlands) and testosterone (Sigma) were added to final concentrations of 2.35 IU/ml and 28 ng/ml (10−7 M) respectively. Both FSH and testosterone concentrations were based on previously published work demonstrating that these doses maximally stimulate the binding of isolated round spermatids to a monolayer of Sertoli cells in vitro (Cameron & Muffly 1991, Perryman et al. 1996). In addition, the dose of FSH used maximally stimulates the in vitro production of two Sertoli cell-derived products, N-cadherin (Perryman et al. 1996) and inhibin (Lampa et al. 1999), and the dose of testosterone used is similar to the levels measured in the 20-day-old rat testis (Killian et al. 2003).
Sertoli cell culture fixation
After culture, Sertoli cell monolayers were fixed with paraformaldehyde (3.7% w/v) in DPB for 10 min at room temperature, followed immediately by permeabilisation with Triton X-100 (Sigma; 0.1% w/v) in DPB for 5 min at room temperature. After fixation, Sertoli cell monolayers were washed twice with cold DPB (4 °C) and stored in DPB at 4 °C until processed for immunocytochemistry. All incubations for immunocytochemistry were performed in a humidified chamber at room temperature, and all solutions, unless otherwise stated, were diluted in 0.01 M PBS, 0.154 M NaCl and 0.1% (w/v) BSA (PBS/BSA).
In vivo studies
In vivo hormone treatments
Testis tissue from rats (75–85 days old) undergoing various hormone treatments were used from a previous study (Meachem et al. 1998), in which 24 rats received 100 μl proprietary GnRH immunogen preparation (BA-1666–4; Biotech, Sydney, Australia) every month for 3 months. This treatment results in the suppression of testicular testosterone levels to 4% of control, serum FSH to close to the limits of detection of the assay (1 ng/ml), and testis weights to 19% of control (Meachem et al. 1998). Control animals received monthly injections of adjuvant alone.
After 3 months of GnRH immunisation, six animals were killed immediately (GnRH-immunised group), and the remaining animals (six per group) received one of the following:
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daily injections of hrFSH (Gonal-F, Serono, Australia; 25 IU/kg per day s.c.) for 7 days (+FSH group)
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administration of 6 cm silastic implants filled with testosterone powder plus daily injections of normal sheep serum (2 mg/kg per day s.c.) for 7 days (+T6 group)
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administration of 6 cm silastic testosterone implants plus daily injections of polyclonal sheep antiserum against FSH (2 mg/kg per day s.c.) for 7 days (+T6+FSH antibody group).
This FSH antibody has been shown to immunoneutralise over 90% of circulating FSH in a normal adult rat and thus prevent the testosterone-dependent restoration of circulating FSH levels in GnRH-immunised animals given testosterone (Meachem et al. 1998). The testis weights, hormone levels and spermatogenic cell populations in these animals are described elsewhere (Meachem et al. 1998). Six additional animals were used for electron microscopy studies: two control animals, two ‘GnRH-immunised group’ animals and two ‘+ FSH group’ animals.
For vinculin immunohistochemical localisation, additional archival testis tissue (Bouin’s fixed) was obtained from a variety of other treatment groups (n=4 per group), as described elsewhere (Meachem et al. 1998); these included GnRH-immunised rats given higher doses of testosterone (24 cm) with or without the FSH antibody for 7 days, control rats given the FSH antibody for 7 days to acutely suppress FSH, and rats given testosterone (3 cm) + 0.4 cm oestradiol (TE) for 9 weeks to suppress serum LH, testicular testosterone (FSH levels are not suppressed) with or without FSH antibody for the last 7 days of treatment.
Tissue fixation
Testes for immunohistochemistry were perfusion fixed in situ with Bouin’s fixative, as described previously (Meachem et al. 1997), and testes for electron microscopic studies were perfused in 5% glutaraldehyde in 0.1 M cacodylate (O’Donnell et al. 2000). After perfusion fixation, Bouin’s-fixed testes were removed, immersed in Bouin’s fixative for approximately 5 h, and then kept in 70% ethanol at 4 °C until use. Testis wedges were embedded in low-melting-temperature ribboning polyester wax (Oke & Suarez-Quian 1993), as described elsewhere (O’Donnell et al. 2000). Sections (7 μm) were cut on a cryostat set at 0 °C, floated onto a water bath set at 32 °C, collected onto slides coated with 2% 3-aminopropyltriethoxy-silane (Sigma), and allowed to dry for 48–72 h at 4 °C. Glutaraldehyde-fixed tissues were prepared for electron microscopy, as described previously (O’Donnell et al. 2000).
Immunocytochemistry
Sertoli cell cultures
Probes for analysis of junctions in vitro were as follows: filamentous actin (phalloidin-TRITC; 10 μg/ml; Sigma), espin (mouse monoclonal IgG; 1:100; BD Transduction Laboratories, Franklin Lakes, NJ, USA), β-catenin (mouse monoclonal IgG; 1:500; BD Transduction Laboratories), vinculin (mouse monoclonal IgG; 1:500; clone hVIN-1; Sigma), pan-cadherin (rabbit polyclonal antibody; 1:500; Sigma), and vimentin (mouse monoclonal IgG; 1:100; clone V9; Sigma). Negative controls consisted of PBS/BSA for phalloidin-TRITC, normal rabbit serum (Dako, Carpinteria, CA, USA) for pan-cadherin, and normal mouse IgG (Serotec, Oxford, UK) for espin, β-catenin, vinculin and vimentin, diluted to concentrations equivalent to the primary probe.
The aim of the current study was to visualise cadherin-containing junctions as a whole rather than to identify the involvement of individual cadherins in inter-Sertoli cell junctions. Given the numerous cadherin family members found in the testis (some 24 cadherin superfamily members were identified in the testis in a recent study (Johnson et al. 2000)), the polyclonal anticadherin antibody used was chosen in order to visualise all classic cadherins at inter-Sertoli cell junctions.
Primary antibodies were detected with either Alexa Fluor 488-goat antimouse IgG (green fluorescence; diluted 1:100; Molecular Probes, Eugene, CA, USA) for espin, β-catenin, vinculin, and vimentin, or Alexa Fluor 568-goat antirabbit IgG (red fluorescence; diluted 1:100; Molecular Probes) for pan-cadherin.
Cover-slips containing immature Sertoli cell monolayers were processed for immunocytochemistry of the above junctional molecules as follows. Cover-slips were cut into eighths with a diamond-tipped pen, and washed twice for 5 min with PBS/BSA. Non-specific binding sites were blocked with CAS Block (Zymed, San Francisco, CA, USA) containing 10% normal sheep serum for 20 min, after which primary antisera in PBS/BSA were added and incubated overnight. After two washes with PBS/BSA at 5 min each, secondary antisera were applied and incubated for 1 h. Finally, cover-slips were washed twice for 5 min with PBS/BSA, counterstained simultaneously with DAPI (4′,6-diamidino-2-phenylindole dihydrochloride; 100 nM; Molecular Probes) and TO-PRO-3 (10 μM; Molecular Probes) for 5 min, rinsed three times with PBS/BSA, and mounted with FluorSave Reagent (Calbiochem, San Diego, CA, USA). For double-label experiments, primary and secondary antisera were incubated on cover-slips simultaneously.
Testis sections
Antibodies for analysis of junctions in vivo were as follows: vinculin (as above, 1:5000), espin (affinity-purified rabbit polyclonal antibody; 1:50 for immunofluorescence; 1:1000 for conventional light immunocytochemistry (Bartles et al. 1996)), actin (mouse monoclonal, 1:250 (clone C4; ICN Biomedicals, Aurora, OH, USA), pan-cadherin (as above, 1:25 000) and vimentin (as above, 1:1000).
Details of immunocytochemical procedures are described elsewhere (O’Donnell et al. 2000). All incubations and dilutions were in PBS (0.01 M PBS, 0.154 M NaCl (pH 7.4) and no sodium azide). Briefly, sections were dewaxed, subjected to antigen retrieval (either microwaved in 0.01 M citrate buffer (pH 6.0) for 10 min or incubated in 0.02% trypsin in 0.1% CaCl2 for 40 min), incubated in 0.3% H2O2, blocked with 300 mM glycine and then 0.1% Triton X-100, and incubated in CAS Block (Zymed, San Francisco, CA, USA) containing 10% normal serum (sheep or rabbit) for 20 min. Sections were incubated with primary antibodies either overnight or for 2 h at room temperature. For conventional light microscopy, sections were incubated for 1 h with an appropriate biotinylated second antibody followed by streptavidin-horseradish peroxidase ABC Complex (Vectastain Elite, Vector Laboratories, Burlingame, CA, USA). The pink-coloured substrate used was Vector VIP (Vector Laboratories), and the sections were counter-stained with Mayer’s haematoxylin (Sigma) prior to dehydration and mounting in DPX, as described previously (O’Donnell et al. 2000). For improved detection of pan-cadherin in the testis, sections were subjected to Tyramide Signal Amplification (TSA kit; NEN Life Science Products, Boston MA, USA) prior to the secondary antibody steps, according to the manufacturer’s instructions. Details of the espin and actin dual-label immunofluorescence procedures and confocal microscopy are given elsewhere (O’Donnell et al. 2000).
Specificity of all primary antibodies was verified by substitution with an equivalent dilution of normal mouse serum (vinculin or actin), preimmune rabbit immunoaffinity-purified IgG (espin), normal rabbit serum (PanCad) or normal mouse IgG (vimentin). In all cases, no significant staining or fluorescence was observed for control antisera in any seminiferous tubules.
Confocal microscopy
Confocal microscopy was performed with a Nikon Diaphot 300 microscope equipped with a krypton/argon laser linked to a BioRad MRC1000 confocal unit (BioRad) and a personal computer with COMOS software (BioRad). Laser lines were at 488, 568, and 594 nm for excitation of Alexa Fluor 488, Alexa Fluor 568 and Phalloidin-TRITC, and TO-PRO-3 respectively. A × 60 oil immersion objective was used, and red, green and far-red channels were scanned sequentially. Colour channels were merged with the Confocal Assistant v 4.02 software package Todd Clark Brelje.
Electron microscopy
Seminiferous tubules were examined at 60 kV with a JEOL 1200EX transmission electron microscope. Areas of inter-Sertoli cell contact were identified under low power (× 3000–7000) and junctional structures were examined at higher power (× 15 000–20 000).
Results
Formation of ES and reorganisation of adherens junction belts is stimulated by FSH, but not testosterone, in immature Sertoli cells in vitro
Confocal analysis of cadherin and β-catenin was used to assess the formation of cadherin-based inter-Sertoli cell AJs in vitro (Fig. 1A–E). As shown in Fig. 1A (forked arrowheads), cadherin and β-catenin colocalised at regions of inter-Sertoli cell contact in the absence of hormones in vitro, suggesting that AJs had formed. These AJs were observed as discrete focal adherens plaques oriented perpendicularly to the cell boundary, and were observed to occur at sites of cytoskeletal actin filament termination (Fig. 1F and G, closed arrowheads). Cadherin, β-catenin and actin localisation in cultures treated with testosterone alone (Fig. 1B and G, arrowheads) did not differ from cultures without hormones (Fig. 1A and F), indicating that this dose of testosterone does not modulate AJ function in immature Sertoli cells. In contrast, the nature and extent of cadherin and β-catenin colocalisation, and therefore AJ formation, was potentiated by FSH and FSH plus testosterone treatment, where AJs were observed as extensive continuous belts around the periphery of these cells (Fig. 1C, D, H and I, arrows).
The colocalisation of cadherin and catenin at sites of inter-Sertoli cell contact could also represent inter-Sertoli cell desmosome-type (intermediate filament-associated) junctions, as there is evidence for (Mulholland et al. 2001) and against (Lee et al. 2004) the association of cadherin with intermediate-filament related junctions in the testis. Therefore, colocalisation of vimentin (intermediate filaments) and cadherin was performed (Fig. 1K–O). In all cultures, staining with vimentin produced a pattern characteristic of intermediate filaments, which were concentrated around the cell nucleus and produced a diffuse network throughout the cell cytoplasm. Occasional intermediate filaments were seen to extend to contact intercellular boundaries perpendicularly; however, colocalisation with cadherin at the cell boundary was not apparent in any treatment (Fig. 1K–N). The intermediate filament staining pattern did not change with FSH and/or testosterone treatment (Fig. 1K–N). The lack of vimentin and cadherin colocalisation indicates that the cadherin-associated adhesion junctions stimulated by FSH contain actin (Fig. 1F–I), but not intermediate filaments.
In the adult rat testis, extensive ES junctions are formed between Sertoli cells and can be assessed by the colocalisation of actin and espin (O’Donnell et al. 2000). Therefore, the colocalisation of these two molecules was assessed in immature Sertoli cells in vitro (see Fig. 1P–T). Although Sertoli cells in vitro expressed actin and espin, these proteins did not colocalise at sites of inter-Sertoli cell contact in the absence of hormones (Fig. 1P, asterisks) or in cultures treated with testosterone alone (Fig. 1Q, asterisks). After treatment with FSH or FSH plus testosterone, extensive colocalisation of actin and espin was observed around the periphery of many cells (Fig. 1R and S, arrows), suggesting ES junctions formed under these conditions. However, not all actin was colocalised with espin, as confocal analysis revealed fibres of both actin and espin oriented parallel to the base of the cells (Fig. 1P–S). Both FSH and FSH plus testosterone treatment appeared to increase the level of espin staining within Sertoli cells (compare Fig. 1P, Q, Z and Z1 with Fig. 1R, S, Z2 and Z3). The pattern of espin and actin staining in cultures treated with testosterone alone did not differ from that in cultures without hormones (compare Fig. 1Q with Fig. 1P).
The colocalisation of espin and cadherin was also investigated to explore the relationship between ES junctions and cadherins (Fig. 1Z–Z4). In the absence of hormones or with testosterone alone, cadherin did not colocalise with espin (Fig. 1Z and Z1, arrowheads) suggesting that unstimulated Sertoli cells form focal AJ puncta, but not ES junctions. In contrast, treatment with FSH or FSH plus testosterone resulted in extensive espin and cadherin colocalisation at the periphery of cells (Fig. 1Z2 and Z3, arrows), suggesting that AJ belts and extensive ESs formed at similar sites between Sertoli cells.
Given that the arrangement of the cytoskeletal molecules actin and vinculin has been shown to be regulated by FSH (Cameron & Muffly 1991, Muffly et al. 1994), the colocalisation of these molecules was also performed (Fig. 1U–Y). In the absence of hormones or with testosterone alone, vinculin was concentrated at the tips of basal actin filaments, suggesting localisation at cell–matrix junctions (Fig. 1U and V, open arrowheads), as well as diffuse localisation within the Sertoli cell cytoplasm (Fig. 1U and V). Upon FSH and FSH plus testosterone stimulation, vinculin became colocalised with actin at the periphery of many cells (Fig. 1W and X, arrows), and the intensity of cytoplasmic vinculin staining appeared to increase (compare Fig. 1W and X with Fig. 1U and V). The localisation of vinculin at the tips of basal actin filaments remained unchanged (Fig. 1W, open arrowheads).
An interesting observation was that regions of the Sertoli cell monolayer that contained a low local density of Sertoli cells (for example, near the edge of the cover-slip, where the Sertoli cell density was approximately 10 Sertoli cell nuclei per 10 000 μm2) formed AJ puncta; however, they did not display the formation of extensive AJ belts or ES junctions (as evidenced by immunostaining for cadherin/catenin and espin/actin), despite stimulation with FSH (data not shown). This suggests that the FSH-stimulated formation of AJ belts and ES junctions is density-dependent.
FSH stimulates the formation of basal ES junctions and the orientation of inter-Sertoli cell junctions in the adult rat testis
The colocalisation of espin and actin by dual-label immunofluorescence confocal microscopy was used to investigate the regulation of inter-Sertoli cell ES junctions in adult testes in vivo. In control animals (where FSH, LH and testosterone levels are normal), espin and actin colocalised in a scalloped pattern parallel to the basement membrane that is characteristic of inter-Sertoli cell ES junctions (Fig. 2A, arrowheads). The colocalisation of espin and actin was observed in all stages of the seminiferous epithelium; however, the nature of this staining differed between stages. During stages I–VIII, a scalloped staining pattern was seen above spermatogonia and early spermatocytes, and below the pachytene spermatocyte-spermatid layers. During stages IX–X, staining was observed both above and below leptotene spermatocytes, while, during stages X–XIV, staining was observed below leptotene and zygotene spermatocytes.
When FSH and LH levels, and thus testicular testosterone levels, were suppressed by long-term immunisation of adult rats against GnRH, limited colocalisation of espin and actin was seen despite the presence of both molecules within the Sertoli cell cytoplasm (Fig. 2B). Espin fibres not colocalised with actin were visible and were arranged in a pattern that was perpendicular to the basement membrane of the tubule (Fig. 2B, arrows). Administration of hrFSH for 7 days to GnRH-immunised animals induced colocalisation of espin and actin into staining patterns characteristic of ES junctions (Fig. 2C, arrowheads). This restoration of ES junctions in Sertoli cells was not apparent when testosterone alone (+T6 + FSH antibody group) was present for 7 days (Fig. 2D), the epithelium having a similar appearance to that of GnRH-immunised animals (Fig. 2B). When both FSH and testosterone were restored in GnRH-immunised animals for 7 days (+T6 group), colocalised espin and actin were seen in a pattern identical to FSH-treated animals (data not shown).
Electron microscopic analysis of ES junctions in these tissues confirmed the findings of the espin/actin colocalisation. Extensive ES junctions were visible between Sertoli cells in normal adult rats (Fig. 3A). This junction is characterised by hexagonally packed actin bundles sandwiched between the Sertoli cell plasma membrane and parallel cisternae of endoplasmic reticulum, which exhibit ribosomes on their membrane closest to the inside of the cell. Within the domain of the ES junction, focal opposing subsurface densities associated with the plasma membrane of each cell were also visible (Fig. 3A, arrows). After GnRH immunisation, extensive ES junctions were not seen between Sertoli cells (Fig. 3B), focal areas of ES junctions being rarely observed. Instead, focal junctions, which appeared morphologically similar to desmosome-like junctions (Vogl et al. 1993), were seen between adjacent Sertoli cells in GnRH-immunised animals (Fig. 3B, arrows). When FSH was given to GnRH-immunised animals, extensive ES junctions, similar to those in Fig. 3A, were again apparent within the epithelium (data not shown).
The immunolocalisation of vinculin, pan-cadherin and vimentin was then examined to compare the localisation of ES, cadherin-based AJs and desmosome-like junctions, respectively, in control, GnRH-immunised and FSH-replaced rat testes. In control rats, vinculin (an ES junction marker) was present in ES junctions between adjacent Sertoli cells (Fig. 2E, arrowheads), in ES junctions between Sertoli cells and spermatids (Fig. 2E, ‘est’), and in the central Sertoli cell cytoplasm (Fig. 2E). Cadherin (PanCad – an AJ marker) was visible at focal locations consistent with staining at inter-Sertoli cell junctions (Fig. 2H, arrowheads, shows an example of cadherin staining at inter-Sertoli cell junctions in the characteristic scalloped pattern). The stage specificity of cadherin staining and its relationship to developing germ cells were identical to that described for the localisation of actin and espin above, although cadherin staining at inter-Sertoli cell junctions was much less extensive than staining of ES junction markers (compare Fig. 2E with 2H). Vimentin (a marker of intermediate filament-related junctions or desmosome-like junctions) was apparent in the perinuclear region and central cytoplasm (Fig. 2K, arrow) of the Sertoli cell in a stage-dependent manner, at sites of Sertoli cell contact with the basement membrane, and around some basally located germ cells, as previously described (Vogl et al. 1993, Zhu et al. 1997). Vimentin staining was also apparent in a scalloped pattern (Fig. 2K, arrowhead) consistent with the known localisation of intermediate filament-related junctions between Sertoli cells (Vogl et al. 1993, Zhu et al. 1997).
In GnRH-immunised animals that lacked FSH and LH, the ordered arrangement of vinculin at ES junctions was lost (Fig. 2F). Cadherin immunostaining (Fig. 2I) was apparent in a linear pattern perpendicular to the basement membrane. Heavy vimentin staining (Fig. 2L) was evident in Sertoli cell cytoplasm and the perinuclear region, near the basement membrane of the tubule and around basally located germ cells.
The organisation of vinculin in ES structures was restored by 7 days of FSH treatment (Fig. 2G, arrowheads), but not by testosterone (6 cm implants), in the absence of FSH (not shown). When FSH was administered to GnRH-immunised rats for 7 days, there was no visible increase in intensity of cadherin or vimentin immunostaining (Fig. 2J and M respectively). However, cadherin immunostaining was again visible in focal regions parallel to the basement membrane in a localisation consistent with inter-Sertoli cell junctions (Fig. 2J). Vimentin immunostaining suggested a more ordered arrangement of intermediate filaments, with a scalloped pattern over basal germ cells more evident (Fig. 2M) compared with GnRH-immunised testes (Fig. 2L). Animals administered 6 cm implants of testosterone plus an FSH antibody (i.e. testosterone in the absence of FSH) showed similar cadherin and vimentin immunostaining to GnRH-immunised animals (data not shown).
The organisation of vinculin at inter-Sertoli cell ES junctions was then investigated in a large number of animals receiving various treatments that manipulate either FSH and/or testosterone (Table 1). The presence of vinculin in the normal scalloped pattern arrangement indicative of ES junctions (Fig. 2E and G), or disorganised vinculin localisation (Fig. 2F) was scored in the various in vivo treated groups. When FSH was present in rats, normal localisation of vinculin at inter-Sertoli cell ES junctions was observed (Table 1). When FSH was acutely withdrawn from control rats for 7 days, normal vinculin localisation was seen. However, when FSH was withdrawn from TE-treated animals, which have low LH and testosterone (but normal ES junctions (O’Donnell et al. 2000)), most animals showed a lack of normal ES pattern. A higher dose of testosterone (24 cm implants) in the absence of FSH could not restore vinculin localisation in ES structures in three out of four rats, although one rat did show normal vinculin localisation. In general, in GnRH-immunised rats, FSH, but not testosterone, could restore normal vinculin localisation, and absence of FSH was associated with absence of normal vinculin localisation.
Discussion
This study investigated the hormonal regulation of inter-Sertoli cell junction formation in vitro and in vivo. Immunostaining for markers of ES junctions, cadherin, catenin and intermediate filaments allowed an appreciation of the formation of several different junction types that are found at sites of inter-Sertoli cell contact, these being ES, AJ, and desmosome-like junctions respectively. In the absence of FSH and testosterone in vitro, immature Sertoli cells were able to form focal AJs, seen as short discrete plaques (AJ puncta), but not ES junctions or desmosome-like junctions. Administration of FSH alone potentiated the formation of extensive junctional plaques containing cadherin, catenin and the ES markers espin, actin and vinculin. In contrast, testosterone alone did not promote extensive junction formation, suggesting that the formation of morphologically mature junctions between immature Sertoli cells in vitro is primarily dependent on FSH. These data suggest that FSH stimulates the formation of ES junctions and extensive AJs between immature Sertoli cells in vitro.
The ability of FSH to stimulate inter-Sertoli cell junction formation, particularly ES formation, was also confirmed in vivo by a GnRH immunisation model. After suppression of FSH, LH and testicular testosterone levels in adult rats for several months, basal ES junctions were disassembled, and ES was absent from the seminiferous epithelium, as assessed by electron microscopy and absence of colocalised actin and espin, which has previously been shown to be an excellent marker of ES junctions in the testis (O’Donnell et al. 2000). Immunostaining for cadherin and vimentin suggested that AJs and intermediate filament-related junctions (desmosome-like junctions) may be present between Sertoli cells; however, their orientation appeared altered. Electron microscopy revealed the presence of desmosome-like junctions between Sertoli cells in GnRH-immunised rats, suggesting that some inter-Sertoli cell junctional types remain during hormone suppression in adult rats, but that ES junctions are completely absent.
Restoration of FSH alone in vivo was able to re-establish ES junctions between Sertoli cells. In addition, immunostaining of cadherin and vimentin suggested that FSH stimulated the correct orientation of AJs and desmosome-like junctions between Sertoli cells, such that the immunostaining pattern was similar to control animals. Taken together, these results suggest that FSH stimulates the formation of extensive inter-Sertoli cell adherens junctions, and that the ES junction in particular is stimulated by FSH.
It is well known that immature Sertoli cells in vitro are highly responsive to FSH; however, it is generally considered that FSH responsiveness declines as Sertoli cells mature (reviewed in Griswold (1993)). Nevertheless, an effect of FSH on inter-Sertoli cell junctions was observed both in immature Sertoli cells in vitro and in adult Sertoli cells in vivo. Previous studies in our laboratory using the experimental in vivo model utilised in the current study identified the importance of FSH in the restoration of spermatogenesis after gonadotrophin suppression (Meachem et al. 1998). Stereological analysis of germ cell numbers showed that long-term gonadotrophin suppression caused arrest of spermatogenesis primarily at the level of spermatogonial and spermatocyte development, and that the restoration of these cell numbers, and subsequently more mature cell types, was dependent on FSH rather than testosterone (Meachem et al. 1998). The current study thus suggests that the re-establishment of mature inter-Sertoli cell junctions by FSH coincides with the restoration of germ cell development (Meachem et al. 1998). Consistent with this, many others have suggested that the formation of mature inter-Sertoli cell junctions is required for spermatogenesis to proceed beyond meiosis, since all post-meiotic cells reside above the blood–testis barrier (Byers et al. 1993).
Although FSH is not required for fertility or the restoration of qualitatively normal spermatogenesis, it is now becoming clear that FSH has an important role in establishing the Sertoli cell population and supporting its function (reviewed in Allan & Handelsman (2005)). Despite the fact that various roles for FSH in the seminiferous epithelium are emerging (see Allan & Handelsman (2005) and O’Donnell et al.(2005)), reports on inter-Sertoli cell junctions in various transgenic models of FSH, LH and androgen action deficiency are limited. However, ultrastructural studies in FSH receptor-deficient (FORKO) mice show various defects in Sertoli cell ultrastructure, particularly fluid-filled cytoplasm indicative of disturbed fluid dynamics (Grover et al. 2004). Although ES structures were observed in FORKO mice, some abnormalities were noted, including dilated endoplasmic reticulum associated with ES (Grover et al. 2004). These studies in transgenic mice support a role for FSH in promoting normal inter-Sertoli cell junctions in vivo.
It was of interest that testosterone was not able to restore ES junctions in the absence of FSH, suggesting that inter-Sertoli cell junctions are particularly FSH sensitive in vivo, as well as in vitro. The observation that AJ and ES junction formation was unaffected by testosterone in both our in vitro and in vivo models was somewhat surprising. The dose of testosterone used in vitro approximates the normal testicular testosterone concentrations in the 20-day-old testis (Killian et al. 2003), and the dose of testosterone used in vivo has been shown to maintain qualitatively normal spermatogenesis (O’Donnell et al. 1999). Although the current and previous (O’Donnell et al. 2000) studies indicate that testosterone is not primarily involved in the regulation of AJs or ES junctions, further work in our (T J Kaitu’u, P Sluka, C F Foo & P G Stanton, personal communication) and other laboratories (Janecki et al. 1991, Florin et al. 2005, Meng et al. 2005) suggests that testosterone regulates the formation of another blood–testis barrier component, inter-Sertoli cell tight junctions, although FSH may also have a role in tight junction permeability, as seen in the hamster (Bergmann 1987, Tarulli et al. 2006) and more recently in the rat (T J Kaitu’u, P Sluka, C F Foo & P G Stanton, personal communication; Janecki et al. (1991)). In addition, it is possible that the normally high levels of testicular testosterone in untreated rats may support the maintenance of AJ and ES junctions to some degree; however, our findings suggest that FSH will be important in the restoration of these junctions after gonadotrophin suppression. We propose that FSH and testosterone cooperate and may even synergise in the initiation and maintenance of inter-Sertoli cell junctions by regulating the various junctional components. It was noted that higher doses of testosterone could potentially mimic the effects of FSH on ES junctions, since one out of four GnRH-immunised rats showed normal vinculin orientation upon treatment with higher-dose testosterone (24 cm implants) plus the FSH antibody. A previous study has also shown that higher doses of testosterone can overcome a lack of FSH in the restoration of germ cell populations (Meachem et al. 1998). Such synergy is commonly noted in the testis with regard to the hormonal regulation of spermatogenesis (reviewed in McLachlan et al. (2002), Meachem et al.(1998), O’Donnell et al.(2005) and Saito et al.(2000)).
The in vitro model used in this study revealed interesting information on the series of events leading to the hormone-dependent formation of AJ and ES junctions between Sertoli cells. AJs are ubiquitous cell–cell adhesion junctions that are characterised by transmembrane cadherin molecules that link to the filamentous actin cytoskeleton via the catenin family of linker proteins (for reviews, see Lui et al. (2003), Nagafuchi (2001) and Yap et al.(1997)). In epithelial cells, AJs are initially formed as AJ puncta at sites of cell–cell contact, found at the tips of interdigitating filopodial extensions. The polymerisation of actin at AJ puncta serves to push adjacent puncta to closer proximity such that they can merge to form extensive AJ belts (Vasioukhin & Fuchs 2001, Vasioukhin et al. 2000). Sertoli cell AJs may also connect to the recently discovered nectin/afadin adhesion complex via related links of these two junction complexes to α-catenin (Takai & Nakanishi 2003). We identified the formation of AJs in Sertoli cells in vitro by the colocalisation of cadherin and β-catenin. In the absence of FSH, AJ puncta were observed, whereas the addition of FSH stimulated the reorganisation of AJ puncta into extensive AJ belts. The formation of both AJ puncta and AJ belts in vitro have previously been demonstrated in the mouse TM4 Sertoli cell line (Sandig et al. 1997) and in mouse keratinocytes (Vasioukhin et al. 2000); however, the progression of AJs in these cells from puncta to belts did not require FSH, probably because these cell types are functionally different from primary Sertoli cell cultures. That the junctions revealed by immunostaining for cadherin and catenin are actin-related AJs, not intermediate filament-related desmosome junctions, is confirmed by the colocalisation of cadherin and actin and the absence of colocalisation of cadherin and vimentin (a component of intermediate filaments) at junction sites.
The specific members of the cadherin superfamily that are found at inter-Sertoli cell AJs remains unknown. The pan-cadherin antibody used in this study is known to detect all classical cadherin forms (E-, N- and P-cadherin) (Geiger et al. 1990), and a similar antibody was used to detect cadherin at inter-Sertoli cell junctions by immunohistochemistry, and to immunoprecipitate β-catenin from seminiferous tubules (Mulholland et al. 2001). More specifically, N-cadherin has been localised to sites of inter-Sertoli cell contact in the adult rat testis (Wine & Chapin 1999, Johnson & Boekelheide 2002, Lee et al. 2003, 2004). Although mRNA for other classic cadherins have been detected in whole adult testis extracts by RT–PCR (Johnson et al. 2000, Lee et al. 2003, Munro & Blaschuk 1996), they have been localised to sites other than inter-Sertoli cell junctions (Byers et al. 1994, Johnson & Boekelheide 2002, Lee et al. 2003, Wine & Chapin 1999). In addition to classical cadherins, a large number of protocadherins and cadherin-related proteins have also been found in testis; however, their localisation has not yet been determined (Johnson et al. 2000). Thus, it is likely that N-cadherin is present at inter-Sertoli cell AJs, but the presence of other cadherin-related proteins cannot be conclusively excluded.
In this study, the ES has been defined as hexagonally packed bundles of actin filaments sandwiched between the Sertoli cell plasma membrane and cisternae of the endoplasmic reticulum. A number of molecular components of ES junctions have been identified, including espin (Bartles et al. 1996) and vinculin (Pfeiffer & Vogl 1991), both of which are codistributed with actin bundles in ES junctions. Espin links actin filaments into hexagonal bundles (Chen et al. 1999, Bartles 2000), while vinculin binds filamentous actin, along with other cell–cell junction components (Menkel et al. 1994, Huttelmaier et al. 1997), serving to link the actin cytoskeleton to the functional adhesion molecules. Our observations on FSH stimulation of actin and vinculin organisation is consistent with data from Muffly and colleagues, who investigated the pattern of actin and vinculin localisation in seminiferous epithelial sheets to suggest that ES junction formation opposite spermiogenic cells is dependent on FSH in vivo (Cameron et al. 1998, Muffly et al. 1993, 1994) and in vitro (Cameron & Muffly 1991).
The fact that ES junctions in vitro were observed only at sites of AJ belt formation, and never at sites of AJ puncta, suggests that the parallel arrangement of actin filaments found at AJ belts (Krendel & Bonder 1999) is important for ES junction formation. It is possible that bundling of actin filaments by espin into the hexagonal arrangement characteristic of ES junctions may occur only after actin filaments are arranged parallel to the plasma membrane. This model of sequential junction formation has been proposed to exist between AJs and tight junctions by Takai and Nakanishi (2003). Once ES junctions are formed, an interdependency of junctions may exist, where the formation of ES junctions may serve to stabilise, and indeed facilitate the organisation of associated cytoskeletal and architectural proteins, thereby strengthening the junctional complex, which includes AJs, and other junctions such as desmosome-like junctions and tight junctions. It was noted that during remodeling of AJ puncta into AJ belts, actin filaments were reorganised to become integral components of AJ belts oriented parallel to the plasma membrane, consistent with current models of actin dynamics (Yonemura et al. 1995, Krendel & Bonder 1999). Although AJs, as defined by cadherin immunostaining, were present between Sertoli cells in adult GnRH immunised rats in vivo, the normal orientation of these junctions appeared dependent on FSH. Thus, it is possible that the formation and orientation of mature inter-Sertoli cell junctions is affected by FSH, possibly via effects on actin reorganisation. Previous studies in adult hypogondotrophic rats support the proposition that FSH is important for the correct organisation of actin in Sertoli cells (Muffly et al. 1993, 1994).
By combining the observations made in this study with data from the literature, we propose a model of FSH stimulation of inter-Sertoli cell junctions in vitro (Fig. 4). In this model, the presence of FSH facilitates the reorganisation of inter-Sertoli cell AJs from puncta to extensive junctions, along with reorientation of the associated actin cytoskeleton and cell membrane. Furthermore, FSH stimulates the recruitment of espin and vinculin, which serve to cross-link actin filaments into the characteristic hexagonal arrangement found in ES junctions. Based on this model, results from the in vivo studies presented here suggest that gonadotrophin suppression causes disassembly of ES junction components, breakdown of AJ belts into AJ puncta, and rearrangement of actin filaments from being arranged parallel to the plasma membrane to terminating perpendicularly at AJ puncta.
In conclusion, we have shown that FSH stimulates the formation of mature inter-Sertoli cell junctions, particularly via effects on the formation of ES junctions and extensive AJs. The effects in immature Sertoli cells in vitro were confirmed in adult testes in vivo, and the restoration of these junctions correlated with the restoration of spermatogenesis. The in vitro results indicate that focal AJs (or AJ puncta), but not ES junctions, form in the absence of hormones, and that FSH promotes the formation of ES junctions and extensive AJ belts. We believe that formation of these junctions is important for mature Sertoli cell function and facilitates the establishment of a functional epithelium capable of supporting spermatogenesis. The in vitro model used in this study provides an interesting model to study the hormone-induced molecular events leading to inter-Sertoli cell junction formation.
Presence (+) or absence (−) of vinculin localisation in ES structures between Sertoli cells in vivo as a function of treatments which result in the presence or absence of FSH (n = 4 animals per treatment*)
| Presence of FSH | Absence of FSH | ||
|---|---|---|---|
| Treatment | Vinculin localin | Treatment | Vinculin localin |
| * Bouin’s-fixed, wax-embedded testis sections were obtained from a previous study (see Meachem et al. (1998) for details). | |||
| a Control animals were administered an FSH antibody for 7 days to suppress acutely FSH in normal animals. | |||
| b Animals given low-dose T (3 cm)+0.4 cm oestradiol (TE) for 9 weeks to suppress serum LH, testicular T; however, FSH levels were not significantly affected (Meachem et al. 1998). | |||
| c Animals given TE for 9 weeks as above; however, FSH antibody was then administered for 1 week to suppress acutely FSH (Meachem et al. 1998). | |||
| d Animals GnRH immunised as described in Materials and Methods, and then given higher dose T (T=24 cm) for the final 7 days treatment, along with FSH antibody to block the concomitant rise in FSH that occurs upon T treatment (Meachem et al. 1998). | |||
| e GnRH+T24 as described above, but with control antibody administered; these animals thus have normal levels of FSH in the final 7 days of treatment (Meachem et al. 1998). | |||
| f One out of four animals showed a vinculin pattern consistent with normal ES; the remaining animals showed disorganized vinculin staining as per GnRH-immunised animals. | |||
| Ab, antibody; T, testosterone. | |||
| Control | + | Control+FSH Abb | + |
| TEb | + | TE+FSH Abc | −/+f |
| GnRH+hrFSH | + | GnRH | − |
| GnRH+T6+control Ab | + | GnRH+T6+FSH Ab | − |
| GnRH+T24+control Abe | + | GnRH+T24+FSH Abd | −/+f |
FSH promotes the formation of adherens junction belts and ES junctions, but not desmosome-like junctions, between Sertoli cells in vitro. Sertoli cells isolated from immature rats were cultured either in the absence of hormones (medium alone), testosterone (28 ng/ml), FSH (2.35 IU/ml) or both FSH and testosterone. Dual-label immunofluorescence was performed and colocalisation analysed by confocal microscopy. Immunostaining was observed as red or green fluorescence, with colocalisation of antigens indicated by yellow staining. Cadherin (red) and β-catenin (green) localisation (A–E) and actin (red) and β-catenin (green) localisation (F–J) were performed to assess the formation of adherens junctions (AJs). Forked arrowheads in (A–D) and (F–H), indicate discrete foci oriented perpendicular to the cell plasma membrane, indicating the formation of discrete AJ puncta; closed arrowheads in (F) and (G), indicate sites of cytoskeletal actin attachment to AJ puncta; arrows in (C) and (D), and (H) and (I), indicate colocalisation of cadherin–catenin and catenin–actin respectively, in extensive zones oriented parallel to the cell plasma membrane, indicating the formation of extensive AJ belts. Vimentin (green) and cadherin (red) localisation (K–O) was performed to assess formation of desmosome-like, or intermediate filament-associated, junctions. Espin (green) and actin (red) localisation (P–T) and vinculin (green) and actin (red) localisation (U–Y) were performed; colocalisation of these molecules (yellow) observed with confocal microscopy suggests the formation of ES junctions. Cultures were also immunostained for espin (green) and cadherin (red) (Z–Z4) to assist with visualisation of cell borders. Arrows in (R) and (S), and (W) and (X), indicate colocalisation of actin–espin and actin–vinculin, respectively, into extensive ES structures at the cell boundary. Asterisks in (P) and (Q), and (U) and (V), indicate absence of ES formation, as evidenced by the absence of espin, actin and vinculin staining at cell boundaries. Open arrowheads in (U–W) indicate colocalisation of vinculin and basal actin filaments associated with cell–matrix, not cell–cell, junctions. Forked arrowheads in (Z) and (Z1), indicate absence of espin and therefore ES junction at discrete cadherin-containing AJ puncta. Arrows in (Z2) and (Z3), indicate colocalisation of cadherin-espin in extensive structures at the cell boundary. Cell nuclei were counterstained with TO-PRO-3 iodide (blue). Negative controls for each immunostaining method are shown in E, J, O, T, Y and Z4. Red staining of cell nuclei is an artefact caused by non-specific binding of the secondary antibody to the cell nucleus. All micrographs were taken at the same magnification, bar = 30 μm. T, testosterone.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
FSH promotes the formation of adherens junction belts and ES junctions, but not desmosome-like junctions, between Sertoli cells in vitro. Sertoli cells isolated from immature rats were cultured either in the absence of hormones (medium alone), testosterone (28 ng/ml), FSH (2.35 IU/ml) or both FSH and testosterone. Dual-label immunofluorescence was performed and colocalisation analysed by confocal microscopy. Immunostaining was observed as red or green fluorescence, with colocalisation of antigens indicated by yellow staining. Cadherin (red) and β-catenin (green) localisation (A–E) and actin (red) and β-catenin (green) localisation (F–J) were performed to assess the formation of adherens junctions (AJs). Forked arrowheads in (A–D) and (F–H), indicate discrete foci oriented perpendicular to the cell plasma membrane, indicating the formation of discrete AJ puncta; closed arrowheads in (F) and (G), indicate sites of cytoskeletal actin attachment to AJ puncta; arrows in (C) and (D), and (H) and (I), indicate colocalisation of cadherin–catenin and catenin–actin respectively, in extensive zones oriented parallel to the cell plasma membrane, indicating the formation of extensive AJ belts. Vimentin (green) and cadherin (red) localisation (K–O) was performed to assess formation of desmosome-like, or intermediate filament-associated, junctions. Espin (green) and actin (red) localisation (P–T) and vinculin (green) and actin (red) localisation (U–Y) were performed; colocalisation of these molecules (yellow) observed with confocal microscopy suggests the formation of ES junctions. Cultures were also immunostained for espin (green) and cadherin (red) (Z–Z4) to assist with visualisation of cell borders. Arrows in (R) and (S), and (W) and (X), indicate colocalisation of actin–espin and actin–vinculin, respectively, into extensive ES structures at the cell boundary. Asterisks in (P) and (Q), and (U) and (V), indicate absence of ES formation, as evidenced by the absence of espin, actin and vinculin staining at cell boundaries. Open arrowheads in (U–W) indicate colocalisation of vinculin and basal actin filaments associated with cell–matrix, not cell–cell, junctions. Forked arrowheads in (Z) and (Z1), indicate absence of espin and therefore ES junction at discrete cadherin-containing AJ puncta. Arrows in (Z2) and (Z3), indicate colocalisation of cadherin-espin in extensive structures at the cell boundary. Cell nuclei were counterstained with TO-PRO-3 iodide (blue). Negative controls for each immunostaining method are shown in E, J, O, T, Y and Z4. Red staining of cell nuclei is an artefact caused by non-specific binding of the secondary antibody to the cell nucleus. All micrographs were taken at the same magnification, bar = 30 μm. T, testosterone.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
FSH promotes the formation of adherens junction belts and ES junctions, but not desmosome-like junctions, between Sertoli cells in vitro. Sertoli cells isolated from immature rats were cultured either in the absence of hormones (medium alone), testosterone (28 ng/ml), FSH (2.35 IU/ml) or both FSH and testosterone. Dual-label immunofluorescence was performed and colocalisation analysed by confocal microscopy. Immunostaining was observed as red or green fluorescence, with colocalisation of antigens indicated by yellow staining. Cadherin (red) and β-catenin (green) localisation (A–E) and actin (red) and β-catenin (green) localisation (F–J) were performed to assess the formation of adherens junctions (AJs). Forked arrowheads in (A–D) and (F–H), indicate discrete foci oriented perpendicular to the cell plasma membrane, indicating the formation of discrete AJ puncta; closed arrowheads in (F) and (G), indicate sites of cytoskeletal actin attachment to AJ puncta; arrows in (C) and (D), and (H) and (I), indicate colocalisation of cadherin–catenin and catenin–actin respectively, in extensive zones oriented parallel to the cell plasma membrane, indicating the formation of extensive AJ belts. Vimentin (green) and cadherin (red) localisation (K–O) was performed to assess formation of desmosome-like, or intermediate filament-associated, junctions. Espin (green) and actin (red) localisation (P–T) and vinculin (green) and actin (red) localisation (U–Y) were performed; colocalisation of these molecules (yellow) observed with confocal microscopy suggests the formation of ES junctions. Cultures were also immunostained for espin (green) and cadherin (red) (Z–Z4) to assist with visualisation of cell borders. Arrows in (R) and (S), and (W) and (X), indicate colocalisation of actin–espin and actin–vinculin, respectively, into extensive ES structures at the cell boundary. Asterisks in (P) and (Q), and (U) and (V), indicate absence of ES formation, as evidenced by the absence of espin, actin and vinculin staining at cell boundaries. Open arrowheads in (U–W) indicate colocalisation of vinculin and basal actin filaments associated with cell–matrix, not cell–cell, junctions. Forked arrowheads in (Z) and (Z1), indicate absence of espin and therefore ES junction at discrete cadherin-containing AJ puncta. Arrows in (Z2) and (Z3), indicate colocalisation of cadherin-espin in extensive structures at the cell boundary. Cell nuclei were counterstained with TO-PRO-3 iodide (blue). Negative controls for each immunostaining method are shown in E, J, O, T, Y and Z4. Red staining of cell nuclei is an artefact caused by non-specific binding of the secondary antibody to the cell nucleus. All micrographs were taken at the same magnification, bar = 30 μm. T, testosterone.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
FSH is involved in the restoration of ectoplasmic specialisation junctions and the organisation of inter-Sertoli cell junctions in vivo. (A, E, H and K) Normal adult rat testis; (B, F, I and L) testis from GnRH-immunised rat; (C, G, J and M) testis from GnRH-immunised rat that was given daily injections of hrFSH for 7 days to restore FSH levels; (D) testis from GnRH-immunised rat that was given 6 cm testosterone-filled implants together with daily injections of an FSH antibody; this treatment partially restores testicular testosterone levels but prevents a concomitant rise in FSH. (A–D) were stained with espin (red) and actin (green) by dual-label immunofluorescence and analysed by confocal microscopy; colocalisation is indicated by the yellow staining. The arrowheads in (A) and (C) show espin and actin colocalised in ES junction structures in a characteristic scalloped pattern. The arrows in (B) and (D) refer to fibres of espin staining that are not colocalised with actin, and that are oriented perpendicular to the basement membrane. (E–G) show vinculin immunostaining as indicated by pink staining. (H–J) show pink immunostaining for cadherin by a broad-specificity (pan-cadherin) rabbit polyclonal antibody that was raised against the cytoplasmic tail common to all classic cadherins. (K–M) show pink immunostaining for vimentin. An example of a negative control slide (in this case rabbit IgG substituted for pan-cadherin antibody) is shown in the inset in (H). (E–M) were counterstained with haematoxylin. Arrowheads in E, G, H, J, K and M indicate immunostaining in a scalloped pattern parallel to the basement membrane, consistent with inter-Sertoli cell junctions; arrows in (I) indicate cadherin staining in an abnormal orientation, being perpendicular to the basement membrane; arrows in (K–M) indicate vimentin staining in Sertoli cell cytoplasm. SC, Sertoli cell nuclei; est, vinculin in ES structures opposite elongated spermatids. The basement membrane of the seminiferous tubule is toward the bottom of each micrograph. Bars = 10 μm; (A–D) were taken at the same magnification, as were (E–M).
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
FSH is involved in the restoration of ectoplasmic specialisation junctions and the organisation of inter-Sertoli cell junctions in vivo. (A, E, H and K) Normal adult rat testis; (B, F, I and L) testis from GnRH-immunised rat; (C, G, J and M) testis from GnRH-immunised rat that was given daily injections of hrFSH for 7 days to restore FSH levels; (D) testis from GnRH-immunised rat that was given 6 cm testosterone-filled implants together with daily injections of an FSH antibody; this treatment partially restores testicular testosterone levels but prevents a concomitant rise in FSH. (A–D) were stained with espin (red) and actin (green) by dual-label immunofluorescence and analysed by confocal microscopy; colocalisation is indicated by the yellow staining. The arrowheads in (A) and (C) show espin and actin colocalised in ES junction structures in a characteristic scalloped pattern. The arrows in (B) and (D) refer to fibres of espin staining that are not colocalised with actin, and that are oriented perpendicular to the basement membrane. (E–G) show vinculin immunostaining as indicated by pink staining. (H–J) show pink immunostaining for cadherin by a broad-specificity (pan-cadherin) rabbit polyclonal antibody that was raised against the cytoplasmic tail common to all classic cadherins. (K–M) show pink immunostaining for vimentin. An example of a negative control slide (in this case rabbit IgG substituted for pan-cadherin antibody) is shown in the inset in (H). (E–M) were counterstained with haematoxylin. Arrowheads in E, G, H, J, K and M indicate immunostaining in a scalloped pattern parallel to the basement membrane, consistent with inter-Sertoli cell junctions; arrows in (I) indicate cadherin staining in an abnormal orientation, being perpendicular to the basement membrane; arrows in (K–M) indicate vimentin staining in Sertoli cell cytoplasm. SC, Sertoli cell nuclei; est, vinculin in ES structures opposite elongated spermatids. The basement membrane of the seminiferous tubule is toward the bottom of each micrograph. Bars = 10 μm; (A–D) were taken at the same magnification, as were (E–M).
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
FSH is involved in the restoration of ectoplasmic specialisation junctions and the organisation of inter-Sertoli cell junctions in vivo. (A, E, H and K) Normal adult rat testis; (B, F, I and L) testis from GnRH-immunised rat; (C, G, J and M) testis from GnRH-immunised rat that was given daily injections of hrFSH for 7 days to restore FSH levels; (D) testis from GnRH-immunised rat that was given 6 cm testosterone-filled implants together with daily injections of an FSH antibody; this treatment partially restores testicular testosterone levels but prevents a concomitant rise in FSH. (A–D) were stained with espin (red) and actin (green) by dual-label immunofluorescence and analysed by confocal microscopy; colocalisation is indicated by the yellow staining. The arrowheads in (A) and (C) show espin and actin colocalised in ES junction structures in a characteristic scalloped pattern. The arrows in (B) and (D) refer to fibres of espin staining that are not colocalised with actin, and that are oriented perpendicular to the basement membrane. (E–G) show vinculin immunostaining as indicated by pink staining. (H–J) show pink immunostaining for cadherin by a broad-specificity (pan-cadherin) rabbit polyclonal antibody that was raised against the cytoplasmic tail common to all classic cadherins. (K–M) show pink immunostaining for vimentin. An example of a negative control slide (in this case rabbit IgG substituted for pan-cadherin antibody) is shown in the inset in (H). (E–M) were counterstained with haematoxylin. Arrowheads in E, G, H, J, K and M indicate immunostaining in a scalloped pattern parallel to the basement membrane, consistent with inter-Sertoli cell junctions; arrows in (I) indicate cadherin staining in an abnormal orientation, being perpendicular to the basement membrane; arrows in (K–M) indicate vimentin staining in Sertoli cell cytoplasm. SC, Sertoli cell nuclei; est, vinculin in ES structures opposite elongated spermatids. The basement membrane of the seminiferous tubule is toward the bottom of each micrograph. Bars = 10 μm; (A–D) were taken at the same magnification, as were (E–M).
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
Electron microscopic analysis of inter-Sertoli cell junctions in vivo. (A) Inter-Sertoli cell junctions in normal adult rat testis. (B) Inter-Sertoli cell junctions, in the absence of ES junctions, in testes from GnRH-immunised rats. In (A) the arrowheads indicate two opposing Sertoli cell plasma membranes, and the arrows indicate two Sertoli cell plasma membranes that are closely associated; focal opposing densities on the plasma membranes are visible. Extensive ES junctions are visible as indicated by actin bundles (a) sandwiched between the Sertoli cell plasma membrane and endoplasmic reticulum (er). Bar = 200 nm. In (B) the arrowheads indicate two opposing Sertoli cell plasma membranes, and the arrows indicate an example of the focal adhesion junctions that were seen. No associated ES junctions were visible. Bar = 500 nm.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
Electron microscopic analysis of inter-Sertoli cell junctions in vivo. (A) Inter-Sertoli cell junctions in normal adult rat testis. (B) Inter-Sertoli cell junctions, in the absence of ES junctions, in testes from GnRH-immunised rats. In (A) the arrowheads indicate two opposing Sertoli cell plasma membranes, and the arrows indicate two Sertoli cell plasma membranes that are closely associated; focal opposing densities on the plasma membranes are visible. Extensive ES junctions are visible as indicated by actin bundles (a) sandwiched between the Sertoli cell plasma membrane and endoplasmic reticulum (er). Bar = 200 nm. In (B) the arrowheads indicate two opposing Sertoli cell plasma membranes, and the arrows indicate an example of the focal adhesion junctions that were seen. No associated ES junctions were visible. Bar = 500 nm.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
Electron microscopic analysis of inter-Sertoli cell junctions in vivo. (A) Inter-Sertoli cell junctions in normal adult rat testis. (B) Inter-Sertoli cell junctions, in the absence of ES junctions, in testes from GnRH-immunised rats. In (A) the arrowheads indicate two opposing Sertoli cell plasma membranes, and the arrows indicate two Sertoli cell plasma membranes that are closely associated; focal opposing densities on the plasma membranes are visible. Extensive ES junctions are visible as indicated by actin bundles (a) sandwiched between the Sertoli cell plasma membrane and endoplasmic reticulum (er). Bar = 200 nm. In (B) the arrowheads indicate two opposing Sertoli cell plasma membranes, and the arrows indicate an example of the focal adhesion junctions that were seen. No associated ES junctions were visible. Bar = 500 nm.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
Model of inter-Sertoli cell junction formation. In the absence of FSH, adjacent Sertoli cells are in contact at interdigitating finger-like projections. At these sites of contact, AJ puncta form, containing cadherin and β-catenin, and are linked to the actin cytoskeleton. Both AJ puncta and actin filaments are oriented perpendicularly to the cell’s plasma membrane. ES junctions do not form in the absence of FSH; however, molecules that are located at ES junctions (espin and vinculin) are seen in the cytoplasm of Sertoli cells. Cell–matrix junctions (hemidesmosomes) containing vinculin linked to basal actin filaments link the Sertoli cell to the basement membrane. After the addition of FSH, actin filament dynamics at Sertoli cell finger-like projections push the plasma membranes of the Sertoli cells together to form extensive sites of inter-Sertoli cell contact, where a junctional complex is formed containing AJ belts, ES junctions and tight junctions. AJ belts form from fused AJ puncta that were once located at adjacent finger-like projections. Actin filaments are now arranged parallel to the Sertoli cell plasma membrane, and are packaged into bundles by espin and vinculin, which move from the cytoplasmic pool into inter-Sertoli cell ES junctions. Linking elements connect adjacent Sertoli cell plasma membranes at sites of ES junction formation, and also connect AJ belts and tight junctions to the actin cytoskeleton. If FSH is removed, ES junctions are disassembled, and the Sertoli cell junctional complex is pushed toward that observed without FSH.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
Model of inter-Sertoli cell junction formation. In the absence of FSH, adjacent Sertoli cells are in contact at interdigitating finger-like projections. At these sites of contact, AJ puncta form, containing cadherin and β-catenin, and are linked to the actin cytoskeleton. Both AJ puncta and actin filaments are oriented perpendicularly to the cell’s plasma membrane. ES junctions do not form in the absence of FSH; however, molecules that are located at ES junctions (espin and vinculin) are seen in the cytoplasm of Sertoli cells. Cell–matrix junctions (hemidesmosomes) containing vinculin linked to basal actin filaments link the Sertoli cell to the basement membrane. After the addition of FSH, actin filament dynamics at Sertoli cell finger-like projections push the plasma membranes of the Sertoli cells together to form extensive sites of inter-Sertoli cell contact, where a junctional complex is formed containing AJ belts, ES junctions and tight junctions. AJ belts form from fused AJ puncta that were once located at adjacent finger-like projections. Actin filaments are now arranged parallel to the Sertoli cell plasma membrane, and are packaged into bundles by espin and vinculin, which move from the cytoplasmic pool into inter-Sertoli cell ES junctions. Linking elements connect adjacent Sertoli cell plasma membranes at sites of ES junction formation, and also connect AJ belts and tight junctions to the actin cytoskeleton. If FSH is removed, ES junctions are disassembled, and the Sertoli cell junctional complex is pushed toward that observed without FSH.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
Model of inter-Sertoli cell junction formation. In the absence of FSH, adjacent Sertoli cells are in contact at interdigitating finger-like projections. At these sites of contact, AJ puncta form, containing cadherin and β-catenin, and are linked to the actin cytoskeleton. Both AJ puncta and actin filaments are oriented perpendicularly to the cell’s plasma membrane. ES junctions do not form in the absence of FSH; however, molecules that are located at ES junctions (espin and vinculin) are seen in the cytoplasm of Sertoli cells. Cell–matrix junctions (hemidesmosomes) containing vinculin linked to basal actin filaments link the Sertoli cell to the basement membrane. After the addition of FSH, actin filament dynamics at Sertoli cell finger-like projections push the plasma membranes of the Sertoli cells together to form extensive sites of inter-Sertoli cell contact, where a junctional complex is formed containing AJ belts, ES junctions and tight junctions. AJ belts form from fused AJ puncta that were once located at adjacent finger-like projections. Actin filaments are now arranged parallel to the Sertoli cell plasma membrane, and are packaged into bundles by espin and vinculin, which move from the cytoplasmic pool into inter-Sertoli cell ES junctions. Linking elements connect adjacent Sertoli cell plasma membranes at sites of ES junction formation, and also connect AJ belts and tight junctions to the actin cytoskeleton. If FSH is removed, ES junctions are disassembled, and the Sertoli cell junctional complex is pushed toward that observed without FSH.
Citation: Journal of Endocrinology 189, 2; 10.1677/joe.1.06634
We thank Dr Sarah Meachem for providing in vivo testis tissue, and Dr David M Robertson for helpful discussions during the course of this work.
Funding This study was supported by Program Grant no. 983212 and 241000 from the National Health and Medical Research Council of Australia (NH&MRC). The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.
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