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6ß1-integrin and phosphorylated focal adhesion kinase between Sertoli cells and elongated spermatids during spermatid release from the seminiferous epithelium
1 Prince Henry’s Institute of Medical Research, Monash Medical Centre, Clayton, Victoria 3168, Australia
2 Department of Anatomy and Cell Biology, Monash University, PO Box 5152, Level 4, Block E, 246 Clayton Road, Clayton, Victoria 3168, Australia
(Requests for offprints should be addressed to L O’Donnell; Email: liza.odonnell{at}princehenrys.org)
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Abstract |
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-integrin form dimerised with ß1-integrin, localising focal adhesion kinase (FAK) and determining if microtubules are involved. Adult Sprague–Dawley rats received testosterone and oestradiol implants and an FSH antibody for 7 days to suppress testicular testosterone and FSH and induce spermiation failure. Control rats were treated with saline. Immunohistochemical analysis showed that 6-integrin and a phosphorylated form of FAK (FAK-Tyr397) are present between late spermatids and Sertoli cells after ES removal, until the point of disengagement, and both proteins remain associated with retained spermatids after spermiation failure induced by hormone suppression. Using dual-label immunofluorescence, tubulins (and thus microtubules) were observed to co-localise with ES, but were neither associated with elongated spermatids just prior to release nor with retained spermatids following hormone suppression. These results suggest that microtubules are not involved in the final release of spermatids from Sertoli cells. We conclude that spermatid release during spermiation is mediated by a ‘disengagement complex’ containing 6ß1-integrin and phospho-FAK, the function of which can be affected by gonadotrophin suppression. |
Introduction |
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Spermiation is vulnerable to disruption via the administration of reproductive toxicants (Huang & Marshall 1983, Chapin et al. 2001) or the suppression of gonadotrophins (Russell & Clermont 1977, Saito et al. 2000, O’Donnell et al. 2001). This results in spermiation failure, wherein spermatids are not released, but instead retained and phagocytosed by Sertoli cells (Russell & Clermont 1977, Russell 1991). The acute suppression of testosterone and FSH in adult rats for 1 week results in 50% of spermatids failing to be released (Saito et al. 2000), suggesting that spermiation failure is an important determinant in the acute onset of spermatogenic suppression after hormone withdrawal. Subsequent studies in monkeys (O’Donnell et al. 2001) and men (McLachlan et al. 2002b) have confirmed this observation.
The molecular mechanisms involved in normal spermiation and spermiation failure are largely unknown. Morphological analysis has showed that spermiation failure induced by gonadotrophin suppression occurs as a result of a defect in the final release or disengagement of spermatids, and not defects in earlier spermiation processes (Beardsley & O’Donnell 2003). In particular, removal of adhesion junctions such as the ES, retraction of the Sertoli cell cytoplasm and removal of spermatid cytoplasm occur normally during gonadotrophin suppression (Beardsley & O’Donnell 2003). Prior to, and during the early part of spermiation, spermatids are attached to the Sertoli cells via the apical Sertoli cell ES, which begin to develop at step 8 of spermiogenesis (Russell 1977, Vogl et al. 2000). Importantly, previous studies showed that this adhesion junction is removed from elongated spermatids ~30 h before disengagement, suggesting that a hitherto uncharacterised adhesion junction is probably involved in spermatid release (Beardsley & O’Donnell 2003). Various cell adhesion molecules and associated kinases are present between spermatids and Sertoli cells during the spermiation process (Chapin et al. 2001); however, the composition of the adhesion complex that mediates spermatid disengagement is unknown.
ß1-integrin is a probable component of this latter junction, as it is present during spermiation, after ES removal and until the point of spermatid release on the dorsal side of the spermatid, and in addition is seen opposite retained spermatids after hormone suppression in adult rats (Beardsley & O’Donnell 2003). 4- (Chapin et al. 2001) and/or 6-integrin (Salanova et al. 1995, 1998) are the probable ß1-integrin partners as both have been immunolocalised to the site of spermiation.
A number of regulatory kinases associate directly or indirectly with ß1-integrin-containing dimers to regulate adhesion processes (Hynes 1992, Sanchez-Mateos et al. 1996, Hannigan & Dedhar 1997). Integrin-linked kinase (ILK) and focal adhesion kinase (FAK) are two kinases that have known effects on ß1-integrin-mediated cell adhesion (Hannigan & Dedhar 1997) and are present in the testis (Mulholland et al. 2001). Despite the demonstration of ILK co-immunoprecipitating with ß1-integrin from the seminiferous epithelium (Mulholland et al. 2001), we recently demonstrated that ILK is associated with spermatids early in spermiation, but is removed along with the ES before disengagement (Beardsley & O’Donnell 2003). Therefore, ILK is not likely involved in spermatid disengagement. Immunolocalisation studies have suggested that FAK is not present at the site of spermiation (Chapin et al. 2001, Mulholland et al. 2001), however, a recent study using antibodies against specific phosphorylated forms of FAK (P-FAK) suggests that P-FAK is indeed present around spermatids during spermiation and immunoprecipitates with ß1-integrin from whole testis extracts (Siu et al. 2003). Given that this kinase is likely to be important in regulating integrin-mediated adhesion (Wehrle-Haller & Imhof 2002, Parsons 2003), it is important to determine whether P-FAK is present at the time of spermatid disengagement and is likely to participate in the regulation of adhesion during spermatid release.
Microtubules are the structural support elements of Sertoli cells and localised mostly within the stalk of the Sertoli cell cytoplasm (Vogl et al. 1993), however, a direct involvement in disengagement is not known. Microtubules, apart from providing structural support, play an important role in the translocation of elongated spermatids from deep within Sertoli cell crypts up to the tubule lumen at the onset of spermiation (Amlani & Vogl 1988, Russell et al. 1989, Redenbach et al. 1992, see Vogl et al. 1993 for review). Microtubules are attached via motor proteins (kinesin and dynein) to the endoplasmic reticulum side of the ES in Sertoli cells (Hall et al. 1992, Miller et al. 1999, Guttman et al. 2000). Immunohistochemical localisation of -tubulin also suggests that microtubules may be present during spermiation (Wenz & Hess 1998), however, whether microtubules are present at the time of spermatid release or are involved in spermatid retention during spermiation failure is unclear.
In order to begin to understand the molecular complexes that control spermatid release, the failure of the disengagement process during gonadotrophin suppression and to elucidate the complexes present on retained spermatids during gonadotrophin suppression, we sought to further characterise the adhesion junction present around spermatids immediately prior to their release. We previously showed that this adhesion junction contains ß1-integrin, but neither ILK nor ES-associated proteins such as espin (Beardsley & O’Donnell 2003). The present study aimed to: (1) identify whether 4- or 6-integrin associates with ß1-integrin at the time of spermatid release, (2) determine if P-FAK (specifically FAK-Tyr397) is part of this ß1-integrin containing complex at disengagement and (3) investigate whether microtubules associate with this complex during spermatid disengagement. The immunolocalisation of these proteins was examined in normal testes, and in testes where spermiation failure had been induced by hormone suppression, in order to pinpoint which adhesion systems may be involved in spermatid retention. We also examined the hormonal regulation of FAK and P-FAK proteins in seminiferous tubules from normal and gonadotrophin-suppressed rats.
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Materials and Methods |
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Sixteen adult male Sprague–Dawley rats (age 70–90 days) were obtained from Monash Central Animal House and housed under 12 h light:12 h darkness photoperiod with access to food and water ad libitum. The study was approved by the Monash Medical Centre Animal Ethics Committee, Australia.
Experimental design
Animals in each of the experiments were assigned into control or treatment groups (n=8 per group). Under isoflurane inhalation anaesthesia, animals in the treatment group received three 8 cm testosterone implants inserted s.c. along the dorsal surface for 1 week to suppress circulating luteinizing hormone, but maintain spermatogenesis as previously described (O’Donnell et al. 1994, Saito et al. 2000). After 7 days, the implants were removed and replaced with a 3 cm testosterone implant and 0.4 cm oestradiol implant (TE treatment) to induce the suppression of testicular testosterone (O’Donnell et al. 1994, 1996). During TE treatment, animals also received daily s.c. injections of rat FSH polyclonal antibody (raised in sheep; 2 mg/kg per day in sterile 0.154 M NaCl), which has been previously shown to immunoneutralise over 90% circulating FSH in adult male rats (Meachem et al. 1998). These animals are referred to as the –testosterone/FSH group. This –testosterone/FSH regime has previously been shown to cause failure of 50% of the spermatids in the testis to spermiate, in the absence of major changes to earlier germ cell populations (Saito et al. 2000). The control group were given daily injections of non-immunised sheep immunoglobulin (ConAb; 2 mg/kg per day in sterile 0.154 M NaCl) for 1 week.
At the end of the treatment regime, four control and four –testosterone/FSH-treated rats were killed by CO2 overdose, one testis was removed, weighed and snap-frozen in liquid N2 for immunohistochemistry. The remaining testis was retained for use in another study. The remaining four control and four treated animals were killed by CO2 overdose and one testis from each rat was removed, weighed and immersion-fixed in Bouin’s fixative for 5 h to be used for morphological analysis and immunohistochemistry. The other testis was removed, weighed and placed into warm PBS (0.01 M PBS, 0.154 M NaCl, pH 7.4, no sodium azide) for seminiferous tubule isolation (see below).
Tissue preparation for immunohistochemistry
For immunohistochemical analysis on frozen tissue, 10 µm sections of frozen testes were cut on a cryostat set at –20 °C and collected onto slides coated with 2%, 3-aminopropyl-triethoxysaline (AAS; Sigma). Sections were then post-fixed in either acetone at –20 °C, or ice-cold Bouin’s fixative for 8 min, rinsed in either cold PBS or cold 100% ethanol respectively, and dried overnight at 4 °C.
Wedges of Bouin’s-fixed testes were embedded in a low-melting-point-ribboning polyester wax (BDH, Poole, Dorset, UK) as previously described (Oke & Suarez-Quian 1993, O’Donnell et al. 2000). Sections of 10 µm were cut on a cryostat set at 0 °C, floated onto a waterbath set at 32 °C, collected onto AAS-coated slides and allowed to dry for 48–72 h at 4 °C.
Immunohistochemistry and double-label immunofluorescence
All antibodies used for immunohistochemistry and Western blot analysis are detailed in Table 1
. The immunohistochemical detection protocol for light microscopy has been described in detail previously (O’Donnell et al. 2000); biotinylated secondary antibodies, a streptavidin–horseradish peroxidase complex (ABC complex; Vectastain Elite, Vector Laboratories, Burlingame, CA, USA) and a pink chromogenic substrate (VIP; Vector) were used to detect the antigen of interest. The signal for FAK-Tyr397 was amplified by employing a tyramide signal-amplification kit (TSA; NEN Life Science products, Boston, MA, USA) as described previously (Beardsley & O’Donnell 2003).
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6-integrin was detected in frozen sections post-fixed in acetone and FAK-Tyr397 was detected in frozen sections post-fixed in Bouin’s fixative. The sections were observed under a 40x objective or a 100x oil-immersion objective on an Olympus BX50 microscope. Images were captured using a FujixHC-2000 high-resolution digital camera (Fujifilm, Tokyo, Japan) and Analytical Imaging Station software (Imaging Research, Inc., St Catherines, ON, Canada). Images were then compiled and labelled using Adobe Photoshop 5.5 (Adobe).
Dual-label immunofluorescence was performed using previously described methods (O’Donnell et al. 2000) and used to co-immunolocalise microtubules and espin. The sections were incubated with the primary antibody overnight at room temperature in a humidified chamber. Following PBS washes, they were incubated in either goat anti-rabbit-546 or goat anti-mouse-488 Alexa fluorophors (Molecular Probes, Eugene, OR, USA) for 1 h in the dark. They were counterstained with Mayer’s hematoxylin (Sigma) and washed in PBS. Coverslips were mounted with Fluorosave (Calbiochem, San Deigo, CA, USA) and were viewed on a confocal microscope (Fluoview FV300, Olympus Australia, Mt Waverley, Vic., Australia). Images from 1 µm optical sections throughout the tissue were collected and compiled using Adobe Photoshop 5.5.
Estimation of the percentage of tubules with elongated spermatids immunostained with adhesion and structural proteins
The estimation of the percentage of tubules during spermiation immunostained with particular antigens is described in detail elsewhere (Beardsley & O’Donnell 2003). Briefly, tubules were scored based on the presence or the absence of spermatids lined along the luminal edge of the tubules, which are termed adluminal spermatids (i.e. beginning of spermiation to disengagement). Tubules with adluminal spermatids were then further classified as to whether spermatids were associated with immunostaining of the antigen of interest (i.e. a positive tubule) or few (<10) or no spermatids with immunostaining (i.e. a negative tubule). The percentage of tubules during spermiation that showed immunostaining associated with the antigen of interest was then calculated. Data were analysed using one-way ANOVA and appropriate post-hoc comparisons using Prism 3.0 software (GraphPad software, San Diego, CA, USA).
Isolation of seminiferous tubules during spermiation
Seminiferous tubules were isolated based on modifications of previously published methods (Chapin et al. 2001). From each of the four control and four –testosterone/FSH animals, one testis was bisected on the longitudinal axis and the contents placed into 20 ml warm digestion media (1 mg/ml collagenase, catalogue no. 103586; 1 mg/ml trypsin, catalogue no. 109819, 0.5 mg/ml hyaluronidase, catalogue no. 106500; Roche; in PBS) and shaken (180 r.p.m.) for 15 min at 32 °C. The enzyme solution was decanted and the tissue rinsed with PBS, followed by PBS+ Mg/Ca. The solution was decanted to ~15 ml and incubated with ~150 kUnitz DNase (DN-25, Sigma) for 3 min at 32 °C with gentle agitation (100 r.p.m.). The tubules were rinsed with PBS, excess PBS was removed by aspiration and seminiferous tubules snap-frozen on dry ice.
Western blot analysis
Protein from whole seminiferous tubules was extracted in a buffer containing 20 mM PO4, 150 mM NaCl, 50 mM NaF, 0.5 mM Na3VO4, 2% v/v Triton-X-100 and complete mini protease inhibitor, EDTA-free (Roche; Chapin et al. 2001). An aliquot was taken at this point to determine the protein concentration using the BCA protein assay (Pierce, Rockford, IL, USA). The supernatant was diluted in a ratio of 1:3 in sample buffer (5 ml 0.5 M Tris–HCl (pH 6.8), 5 ml glycerol, 5 ml 10% w/v SDS, 0.5 ml ß-mercaptoethanol and 24.5 ml MilliQ water). Twenty micrograms protein per sample were separated by SDS-PAGE (7.5% agarose) and transferred onto nitrocellulose membranes for Western blotting. The membranes were blocked in 5% skim milk in PBS overnight at 4 °C. FAK-Tyr397 was detected using an Infrared 680-conjugated goat anti-rabbit secondary antibody (catalogue no. A21076 [GenBank] ; Molecular Probes). FAK and actin were detected using an Infrared 800-conjugated goat anti-mouse secondary antibody (catalogue no. 610132121; Rockland Immunochemicals, Inc., Gilbertsville, PA, USA). The membranes were scanned on an infrared imaging system (Odyssey IR imaging scanner; Li-Cor Biosciences, Lincon, NE, USA) at a resolution of 169 µm and a laser intensity of 5. Results were quantitated using densitometry software provided with the imaging system and data normalised to actin.
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Results |
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Integrins
At the beginning of spermiation, when elongated spermatids were lined along the luminal edge (stage VII), intense 6-integrin immunostaining was seen around the heads of step 19-elongated spermatids (Fig. 1A). The localisation of 6-integrin around these spermatids was concentrated around the dorsal aspect of their heads (Fig. 1A and B) and persisted until the spermatids were released from Sertoli cells during mid-stage VIII, since all stage VIII tubules with elongated spermatids showed 6-integrin immunostaining. Immunostaining was also seen in focal points of developing ES junctions opposite step 8 round spermatids (Fig. 1B). However, obvious immunostaining was not observed in association with steps 9–18 elongating and elongated spermatids (data not shown). After spermatid disengagement, 6-integrin was not seen along the luminal edge of late stage VIII–early IX tubules (not shown). 6-integrin immunostaining was occasionally very faintly associated within the basal compartment consistent with an inter-Sertoli cell junction localisation (not shown). In –testosterone/FSH animals, the localisation of 6-integrin was identical to that seen in control animals in all the tubules examined, i.e. immunostaining was seen until the point of spermatid disengagement. 6-integrin was also seen around many retained spermatids within the epithelium (Fig. 1C), and particularly prominent around retained spermatids in the adluminal portion of the seminiferous epithelium. Staining intensity was reduced and eventually not seen as spermatids were translocated down to the basement membrane and digested by the Sertoli cell (Fig. 1C). The localisation of 6-integrin mirrors previous reports seen for ß1-integrin (Beardsley & O’Donnell 2003).
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4-integrin could not be detected using four different antisera (see Table 1
) and a variety of fixation and detection protocols. In addition, a specific signal was not observed in seminiferous tubule lysates by Western blotting using the same antibodies (not shown). Thus, we cannot make any assessment of 4-integrin reactivity in the seminiferous epithelium.
FAK-Tyr397
Prior to spermiation in stages IV–VI, FAK-Tyr397 localisation was seen in discrete focal bands associated with clusters of steps 17–18-elongated spermatids within Sertoli cell crypts (data not shown). During spermiation, FAK-Tyr397 was localised around the dorsal curvature of step 19-elongated spermatids in stages VII and VIII (Fig. 2A and B), consistent with previous reports (Siu et al. 2003). This localisation persisted until they were released, as all tubules in stage VIII with step 19 spermatids showed FAK-Tyr397 localisation to elongated spermatids (Fig. 2B) similar to the localisation of 6-integrin. FAK-Tyr397 was also present in developing ES opposite steps 8 and 9 spermatids in both control and –testosterone/FSH rats (Fig. 2B and C). In –testosterone/FSH rats, the localisation of FAK-Tyr397 was observed until spermatid release as seen in control animals. Furthermore, FAK-Tyr397 immunostaining was seen on many retained spermatids in the adluminal portion of the epithelium (Fig. 2C).
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Microtubules
The tubulins (both and ß) were immunolocalised in the seminiferous epithelium during spermiation, most prominently within the central stalk of the Sertoli cell cytoplasm (Fig. 3A). They were also evident in the spermatid flagella that were in the tubule lumen throughout the spermiation process and until spermatids were released (not shown). They were also apparent around the heads of elongated spermatids at the beginning of spermiation in stage VII (Fig. 3A). During stage VIII, however, tubulins became more diffusely associated with the heads of elongated spermatids and, immediately prior to spermatid disengagement, there was no immunolocalisation of tubulins associated with spermatid heads (Fig. 3B). Instead, tubulin staining was visible in central Sertoli cell cytoplasm and step 8 round spermatid manchettes (opposite developing ES junctions; Fig. 3B). In –testosterone/FSH rats, the immunolocalisation pattern of tubulins was identical to that seen in control testes. In addition, tubulins did not appear to be immunolocalised around retained spermatids (Fig. 3C), but rather were located within the Sertoli cell cytoplasm.
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). In stage VIII tubules (just prior to spermatid disengagement), espin was no longer associated with the heads of elongated spermatids. While tubulins did not co-localise with espin during this stage, they were also not associated with elongated spermatids and rather localised to the round spermatids manchettes and within the Sertoli cell cytoplasm (Fig. 4E–H). In –testosterone/FSH rats, the same immunolocalisation pattern was seen as in control animals until disengagement, and neither tubulin nor espin were associated with retained spermatids within the epithelium (Fig. 4I–L).
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The percentage of spermiation tubules with immunolabelled spermatids was determined in order to compare the time in which the proteins of interest were present during spermiation (Beardsley & O’Donnell 2003). During spermiation, 6-integrin was associated with spermatids in 94 ± 1.05% (mean ± S.E.M.) of tubules (see Table 2); this is comparable to that seen previously for ß1-integrin (93 ± 0.55% of tubules (Beardsley & O’Donnell 2003)). FAK-Tyr397 associated with spermatids in 92 ± 0.25% of tubules and is comparable to the integrins. Both integrins and FAK-Tyr397 remained associated with spermatids for a longer period of time than the ES protein espin (P<0.05 compared with 6-, ß1-integrin and FAK-Tyr397). It was noted that unlabelled tubules probed for 6-integrin and FAK-Tyr397 were seen during early (stage VII) rather than late spermiation prior to disengagement, whereas tubules that were negative for espin were noted during late spermiation (stage VIII).
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- and ß-tubulin were immunolocalised to spermatid heads in 80 ± 1.32% and 76 ± 1.04% of tubules respectively (Table 2). It was noted that the majority of unlabeled tubules (20 and 24% for - and ß-tubulin respectively) were seen during stage VIII. Tubulins remained associated with step 19-elongated spermatid heads for the same time as espin as there was no significant difference between the percentage of espin and /ß-tubulin-positive tubules. Western blot analysis
Since FAK-tyr397 immunolocalisation remained associated with retained spermatids in stage IX seminiferous tubules from –testosterone/FSH rats, it was hypothesised that FAK phosphorylation may be hormone regulated. Both phosphorylated (FAK-Tyr397) and FAK were detected at the expected molecular mass of 125 kDa in seminiferous tubules from control rats and in tubules from –testosterone/FSH rats. Comparative Western blots and densitometric analysis revealed that there was no significant change in either FAK-Tyr397 or FAK protein levels in seminiferous tubules during spermiation failure compared with controls (Fig. 5).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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6-integrin and FAK-Tyr397. The demonstration of these proteins between Sertoli cells and elongated spermatids until the time of disengagement, and on retained spermatids during spermiation failure, suggests that they play a key role in the release of spermatids during spermiation. These results extend previous findings in which ß1-integrin also forms part of this complex (Beardsley & O’Donnell 2003). Thus, spermatid disengagement, and in particular the failure of this process during gonadotrophin suppression, may be hormonally regulated via an integrin/phosphorylated FAK-containing complex. Furthermore, we show that microtubules associate neither with this complex, nor with retained spermatids after phagocytosis by the Sertoli cell. We conclude that spermatid release at the end of spermiation is mediated by a ‘disengagement complex’ containing 6ß1-integrin and phosphorylated FAK. While the ES junction is well known to mediate adhesion between spermatids and Sertoli cells, this junction is removed prior to the final disengagement of spermatids at the end of spermiation (Beardsley & O’Donnell 2003). Previous studies have proposed various molecules and molecular pathways at the spermatid–Sertoli cell junctional complexes during the entire spermiation process occurring in stages VII and VIII of the rat spermatogenic cycle (Wine & Chapin 1999, Chapin et al. 2001, Mulholland et al. 2001) and reviewed by Lui et al. (2003b), Lee & Cheng (2004), Siu & Cheng (2004), Wong & Cheng (2005) and Zhang et al.(2005); however, little effort has focused on identifying the molecules and processes involved specifically after the removal of ES. In addition, there are limited studies investigating the hormonal regulation of spermiation and disengagement, reviewed by Russell (1991) and McLachlan et al. (2002a). Spermatid release is the first process to show morphological signs of damage after hormone suppression in rodents (Russell & Clermont 1977, Saito et al. 2000) and failure of this process contributes to acute and chronic sperm count suppression in contraceptive-treated monkeys and men (Zhengwei et al. 1998, O’Donnell et al. 2001, McLachlan et al. 2002b).
Previously, we and other researchers have shown that ß1-integrin is present during spermiation at the Sertoli cell–spermatid junction (Palombi et al. 1992, Salanova et al. 1995, Chapin et al. 2001, Mulholland et al. 2001, Beardsley & O’Donnell 2003, Siu et al. 2003). Furthermore, ß1-integrin persists along the dorsal curvature of the spermatid (which is the site where the Sertoli cell remains in contact with the spermatid after it has retracted away from the rest of the cell), until the point of disengagement and present on retained spermatids during spermiation failure (Beardsley & O’Donnell 2003). Both 4- and 6-integrin have been immunolocalised to the spermatid–Sertoli cell junction during spermiation (Salanova et al. 1995, Chapin et al. 2001); however, the persistence of these proteins until disengagement and their presence on retained spermatids had not been investigated. Despite previous reports of 4-integrin in the testes (Chapin et al. 2001), we could not detect 4-integrin immunoreactivity using multiple antisera; thus, we cannot further assess the role of 4-integrin in spermiation.
Although the localisation of 6-integrin to the site of spermiation has been previously demonstrated (Salanova et al. 1995, Chapin et al. 2001), the present study extends previous findings to show that this integrin, along with ß1-integrin, persists after the ES has been removed and until disengagement, indicating that 6ß1-integrins are part of the disengagement complex that mediates spermatid release. The fact that both 6 and ß1-integrin associate with newly retained spermatids after hormone suppression-induced spermiation failure supports this proposition.
The molecular mechanisms regulating the adhesive function of the 6ß1-integrin dimer during spermiation are unknown. Various adhesion-related signalling molecules have been either immunolocalised to the site of spermiation or co-immunoprecipitated with integrins and other ES-related molecules during the spermiation process (Chapin et al. 2001, Siu et al. 2003, Lui et al. 2003a, Lee et al. 2005). In particular, protein phosphorylation seems to be important in spermatid disengagement (Chapin et al. 2001) and thus, signalling cascades are presumably involved in regulating the function of the 6ß1-integrin dimer. Studies using an in vitro culture of spermiation showed that spermatid release decreased when serine/threonine kinase activity was inhibited, yet was increased with the addition of okadaic acid, a serine/threonine phosphatase inhibitor (Chapin et al. 2001), supporting the concept that spermatid release is controlled by protein phosphorylation. Consistent with this, antibodies raised against phosphorylated serine, threonine and tyrosine proteins immunostain the Sertoli cell–spermatid junction during spermiation (Chapin et al. 2001, Mulholland et al. 2001).
Focal adhesion kinase is a key regulator of integrin-mediated adhesion, reviewed by Giancotti & Ruoslahti (1999). Previously, FAK was reported not at the site of spermiation, but rather towards the basal aspect of the epithelium (Mulholland et al. 2001). More recent studies confirmed this, but also showed that phosphorylated FAK forms Tyr397 and Tyr576 were present between spermatids and Sertoli cells during spermiation, leading the authors to conclude that phosphorylated FAK is a component of the ES (Siu et al. 2003). FAK is auto-phosphorylated on its tyrosine397 residue following an association with the cytoplasmic tail of ß1-integrin (Giancotti & Ruoslahti 1999). The present study shows that FAK-Tyr397 co-localises with 6ß1-integrin not only at the site of the ES between spermatids and Sertoli cells early in spermiation, but that this complex persists after ES removal until spermatid disengagement. Although it has been previously demonstrated that FAK-Tyr397 is present in ES junctions (Siu et al. 2003) and likely to be involved in junction formation and turnover (Wong & Cheng 2005), the present data strongly suggest that FAK-Tyr397 is present in association with spermatids immediately before they are released, and therefore not solely a part of ES junctions. Moreover, the demonstration of FAK-Tyr397 immunostaining on retained spermatids during spermiation failure supports the hypothesis that spermatid release and retention involves an 6ß1-integrin/FAK-Tyr397 complex.
De-phosphorylation of FAK on the tyrosine397 residue is associated with FAK-exiting adhesion sites and the subsequent disassembly of integrin-mediated adhesion complexes (Kabir et al. 2002). It is therefore plausible to speculate that FAK de-phosphorylation is a key mediator of spermatid disengagement. The observation that FAK-Tyr397 associated with retained spermatids during spermiation failure supports this hypothesis. In order to investigate changes in FAK phosphorylation, protein levels of FAK and FAK-Tyr397 were investigated in seminiferous tubules. Given the persistence of FAK-Tyr397 on retained spermatids, we expected to see an increase in FAK-Tyr397 protein in seminiferous tubules from gonadotrophin-suppressed rats. The fact that no changes were seen may reflect that changes in FAK-Tyr397 at other stages of the cycle may mask any change after disengagement, or an induction of a 50% spermiation failure rate is not enough to see any changes in the overall protein levels.
Results from this study, together with our previous study (Beardsley & O’Donnell 2003), suggest that spermatid disengagement and the retention of spermatids after gonadotrophin suppression is mediated by an adhesion complex containing 6ß1-integrin and FAK-Tyr397. This complex is separate from the ES, and thus we use the term ‘disengagement complex’ to distinguish between the adhesion complex present between Sertoli cells and spermatids at the time of disengagement, from the better-known ES junction, which is present between Sertoli cells and spermatids from mid-spermiogenesis until mid-spermiation (Fig. 6). While the ES has a distinct ultrastructure, reviewed by Vogl et al.(2000), Mruk & Cheng (2004), the disengagement complex between spermatids and Sertoli cells just prior to the release lacks ultrastructural features (Beardsley & O’Donnell 2003). However, molecularly, the disengagement complex resembles a focal adhesion complex (FAC), which is a well known adhesion system involved in cell migration and anchorage (Martin et al. 2002, Wehrle-Haller & Imhof 2002). Integrin-containing FACs are dynamic junctions involved in rapid adhesion and loss-of-adhesion events. That the disengagement complex is a type of FAC is supported by the localisation of 6ß1-integrin and phosphorylated FAK, as well as the localisation of other FAC-associated proteins to the site of spermiation, for example, paxillin, cortactin, Ras, RalA and Rac1 (Wine & Chapin 1999, Chapin et al. 2001). This disengagement complex is likely assembled at the beginning of spermiation, when the 6ß1-integrin molecules cluster on the Sertoli cell plasma membrane opposite the dorsal curvature of the step 19 spermatid, and persists after ES removal (Beardsley & O’Donnell 2003 and present study). The degree of integrin clustering in well-developed FACs is proportional to junctional strength (Giancotti & Ruoslahti 1999, Shemesh et al. 2005) and the heavy integrin immunostaining around the dorsal curvature of the spermatid would suggest that the disengagement complex has a strong adhesive function. Consistent with this hypothesis is the fact that, for at least a day prior to release, elongated spermatids are extended well out into the tubule lumen and thus presumably encountering considerable shear forces from seminiferous tubule fluid flow, with the Sertoli cell only contacting its the dorsal curvature (Russell 1993, Beardsley & O’Donnell 2003). The ES, which is well known to have considerable adhesive strength (Vogl et al. 2000, Wolski et al. 2005), is not present at this time (Beardsley & O’Donnell 2003) and thus the disengagement complex must be of sufficient strength to prevent premature sperm release. Furthermore, the demonstration of the disengagement complex on retained spermatids suggests that it is the dysfunction of this complex during gonadotrophin suppression that underlies spermiation failure.
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Lastly, this study investigated whether the Sertoli cell microtubule network was involved in sperm release. It is well known that microtubules have a structural role as well as a transport role within the Sertoli cell (Russell et al. 1989, Vogl et al. 1993). Microtubules attach to the Sertoli cell ES and assist in the translocation of elongated spermatids prior to spermiation (Amlani & Vogl 1988, Russell et al. 1989, Vogl et al. 1993, Guttman et al. 2000) and are involved in the extension of the Sertoli cell stalk into the tubule lumen during spermiation (Russell 1993). Given their translocation and structural functions within the seminiferous epithelium, we hypothesised that microtubules may be involved in spermatid release and/or retention after hormone suppression. The immunolocalisation of tubulins, co-localised with the ES protein espin, was employed to map the localisation of tubulins during spermiation. The results presented here suggest that microtubules, while assisting in spermatid translocation, do not appear to play a direct role in spermatid release, since tubulins did not associate with spermatids at the time of disengagement. Likewise, microtubules did not appear to be involved in the initial retention of spermatids during spermiation failure. However, given their abundance and role in transport within the Sertoli cell, reviewed in Vogl et al.(1993), they may well be involved in the translocation of the retained spermatids down to the basement membrane after they have been phagocytosed by the Sertoli cell. We did not see any difference in the immunolocalisation patterns between controls and gonadotrophin-suppressed seminiferous tubules prior to, or after spermatid release, suggesting that microtubule localisation is not hormonally regulated.
In summary, this study provides further insights into mechanisms of spermatid release during spermiation and the retention of spermatids after gonadotrophin suppression-induced spermiation failure. The microtubule network, while may be providing structural and movement roles, does not appear to be directly involved in the disengagement or in the retention of spermatids. However, a disengagement complex containing 6ß1-integrin and FAK-Tyr397 is present between Sertoli cells and elongated spermatids during spermiation until spermatids are released from the epithelium. The results support the contention that a defect in the function of the disengagement complex is involved in spermatid retention during gonadotrophin suppression.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Received 15 March 2006
Received in final form 8 May 2006
Accepted 20 June 2006
Made available online as an Accepted Preprint 29 June 2006
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