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Testis |
a Prince Henry's Institute of Medical Research, Clayton, Victoria 3168, Australia
b Department of Anatomy and Cell Biology, Monash University, Clayton, Victoria 3168, Australia
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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follicle-stimulating hormone, Sertoli cells, spermatid, spermatogenesis, testosterone
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The control of spermiation and spermatid release is not understood; however, it has been suggested that cell adhesion molecules and associated kinases assist in adhesion of spermatids to Sertoli cells during spermiation and, thus, may control sperm release [3, 4]. Elongated spermatids are attached to Sertoli cells via the ES [5–7]; however, this strong adhesion junction seems to be removed before disengagement of the spermatid [2]. Thus, how spermatids adhere to Sertoli cells just before disengagement is not known. ß1-Integrin [8–10] and one of its associated kinases, integrin-linked kinase (ILK) [4, 11], are candidates for mediating adhesion between the spermatid and Sertoli cell during spermiation, as are N-cadherin and desmoglein, as evidenced by immunohistochemistry [12]. A recent study proposed a model of a multiprotein complex responsible for sperm release [4], suggesting a complex series of protein interactions during spermiation. However to our knowledge, the actual molecules responsible for sperm release have not been identified.
Spermiation is an important determinant of sperm output from the testis and is a vulnerable process that can be easily disrupted. Various treatments, including reproductive toxicants [13, 14] and gonadotropin suppression [15, 16], result in spermiation failure, in which spermatids are not released but instead are retained and phagocytosed by the Sertoli cell [2, 17, 18]. Quantitative stereological methods used in our previous study [18] showed that after 1 wk of testosterone (T) and FSH suppression in rats, 50% of the spermatids in the testis failed to spermiate and were retained, suggesting that spermiation failure is a major factor in the acute onset of spermatogenic suppression after hormone withdrawal. Subsequent studies in monkeys [19] and humans [20] showed that spermiation failure also occurs after gonadotropin suppression induced by androgen-based contraceptive administration. Importantly, the latter study [20] demonstrated that the induction of spermiation failure in humans is important for the acute and chronic suppression of sperm counts during contraceptive administration and is a key determinant of contraceptive effectiveness.
Despite the clinical importance of spermiation failure, relatively little is known about the events associated with spermiation failure after hormone suppression. Electron-microscopic observations in hypophysectomized rats suggested that spermiation failure might involve the abnormal retraction of the Sertoli cell cytoplasm and the failure of TBCs to form correctly [16]. However, more detailed studies regarding the morphological and ultrastructural events associated with spermiation failure induced by hormone suppression are required.
The present study aimed to determine the effects of acute hormone suppression on the morphological and ultrastructural events associated with spermiation and to determine the localization of cell adhesion (integrins and cadherins) and associated (ß-catenin and ILK) molecules during hormone suppression-induced spermiation failure. A model of T and FSH suppression for 1 wk was used to induce spermiation failure in at least half the spermatids in the testis, as previously described [18], to define key processes and to investigate the localization of putative molecules involved in spermiation failure.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Eight adult male Sprague-Dawley rats (age, 70–90 days) were obtained from Monash Central Animal House and housed under a 12L:12D photoperiod with free access to food and water. The study was approved by the Monash Medical Center Animal Ethics Committee.
Experimental design
Eight- and 3-cm implants filled with T powder (Sigma, St. Louis, MO) and 0.4-cm implants filled with estradiol powder (Sigma) were prepared as described previously [21]. Four animals in the T and FSH suppression (-T&FSH) treatment group were anesthetized by ether inhalation, and implants were placed s.c. along the dorsal surface. Animals initially received three 8-cm implants (T24 implants) for 1 wk to suppress circulating LH but maintain spermatogenesis as previously described [18, 21]. After 1 wk, the T24 implants were removed and replaced with 3-cm T and 0.4-cm estradiol implants (TE treatment) for 1 wk to induce suppression of testicular T [21, 22]. During TE treatment, animals also received daily s.c. injections of rat FSH polyclonal antibody (raised in sheep) at a concentration of 2 mg kg-1 day-1 in sterile 0.154 M NaCl, which has been previously shown to immunoneutralize more than 90% of circulating FSH in adult male rats [23]; these animals formed the -T&FSH group. This -T&FSH regime has been previously shown to have a synergistic effect on spermiation failure, causing a more marked suppression than either T or FSH alone, and to have caused 49.7% ± 5% (mean ± SEM, n = 6) of spermatids to fail to spermiate [18]. The remaining four animals in the present study did not receive any implants but were given daily injections of nonimmunized sheep immunoglobulin at a concentration of 2 mg kg-1 day-1 in sterile 0.154 M NaCl for 1 wk and, thus, formed the control group. This regime has been previously shown to have no quantitative effects on spermiation [18].
Tissue Preparation
At the end of the experiment (on the eighth day of hormone suppression), four control and four -T&FSH rats were anesthetized by ether. One testis was excised, weighed, and frozen in isopentane cooled in liquid nitrogen and then stored at -80°C for subsequent immunohistochemical analysis. The remaining testis was perfusion-fixed with 5% glutaraldehyde in 0.1% cacodylate buffer for light- and electron-microscopic analysis as previously described [24]. After postfixation with 5% glutaraldehyde in 0.1% cacodylate buffer for 24 h at 4°C, the perfusion-fixed testes were sliced in half. For light-microscopic analysis, one half was cut into 2-mm slices along the long axis of the testes. From these, 3- x 2-mm blocks were processed into Epon araldite, and 1-µm sections were cut from each block using glass Ralph knives on a Supercut microtome (model 2050; Reichert Jung, Noffloch, Germany). Sections were air-dried on slides overnight at 37°C, stained with toluidine blue, and mounted with DPX (BDH, Poole, U.K.).
For electron-microscopic analysis, the second half of the perfused testes was prepared as previously described [24]. For immunohistochemical analysis, 10-µm sections of frozen testes were cut on a cryostat set at -20°C and melted onto slides coated with 2% 3-aminopropyl-triethoxy-saline (AAS; Sigma). Sections were then postfixed in acetone at -20°C for 8 min, rinsed in cold PBS (0.01 M phosphate buffer and 0.154 M NaCl, pH 7.4, no sodium azide) and dried overnight at 4°C. For some immunohistochemical analyses, Bouin-fixed, wax-embedded testes were used from control and -T&FSH rats from a previous study [18]. Wedges of testis were embedded in a low-melting-point ribboning polyester wax (BDH) [24, 25]. Ten-micrometer sections 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.
Electron Microscopy
Each section was examined at 60 kV using a JEOL 1200EX transmission electron microscope (Jeol Australasia Pty. Ltd., Brooksvale, Australia). Each seminiferous tubule was examined on low power (3000–7000x) for identification of spermatids during spermiation. Following this identification, spermatids were examined on higher power (
15 000–30 000x). Tubules were classified into the following stages for examination of spermiation: early stage VII, midstage VII, late stage VII, transition between stages VII and VIII, early stage VIII, late stage VIII, and stage IX [26]. One or two sections per animal (n = 4 per group) were observed, and each section contained one or two tubules during spermiation. An average of 9 or 10 spermatids was evaluated per tubule.
Light Microscopy
One-micrometer sections stained with toluidine blue were viewed on an Olympus BX50 microscope (Olympus, Melbourne, Australia), and images were captured using a FujixHC-2000 high-resolution digital camera (Fujifilm, Tokyo, Japan) and Analytical Imaging Station software (Imaging Research, Inc., St. Catharines, ON, Canada). Images were then compiled and labeled using Adobe Photoshop 5.5 (Adobe, Palo Alto, CA). Specifically, stages V–IX were examined, viewing approximately 10 tubules per animal (n = 4 per group).
Immunohistochemistry and Double-Label Immunofluorescence
ß1-Integrin was detected using a rabbit polyclonal antiserum raised against a synthetic polypeptide corresponding to the C-terminal cytoplasmic domain of human ß1-integrin (catalog no. AB1952; Chemicon, CA, USA) at a dilution of 1:500. The negative control used was an equivalent dilution of normal rabbit serum (DAKO, Carpinteria, CA). The ILK was detected using an immunoaffinity-purified polyclonal antiserum raised against a GST fusion protein containing the kinase domain of human ILK (catalog no. 06-592; Upstate Biotechnology, Lake Placid, NY) at a dilution of 1:100. The negative control used was an equivalent dilution of rabbit immunoglobulin (Ig) G (DAKO).
To detect cadherins, a rabbit polyclonal antibody raised against the C-terminal amino acids of chicken N-cadherin (catalog no. C-3678; Sigma) was used at a dilution of 1:25 000. This region is highly conserved between members of the cadherin family; thus, this antibody detects all forms of classic cadherins and is referred to as a Pan-cadherin (PanCad) antibody. The negative control used was an equivalent dilution of normal rabbit serum (DAKO). To detect N-cadherin during spermiation, Sigma clone GC-4, Sigma clone FA-5, and Santa Cruz N-19 (catalog no. SC-1502; Santa Cruz Biotechnologies, Santa Cruz, CA) antibodies were used at a number of dilutions and in various fixed tissues, including Bouin, formalin, and methacarn, and in unfixed frozen tissue. No specific staining was observed above background level using Sigma clones GC-4 and FA-5; thus, only Santa Cruz N-19 was used for analysis. An immunoaffinity-purified polyclonal antibody, Santa Cruz N-19 was used at a dilution of 1:500. The negative control used was an equivalent dilution of purified mouse IgG (DAKO). ß-Catenin was detected with a rabbit polyclonal antibody raised against the N-terminus of ß-catenin (kindly provided by Dr. Alpha Yap, Department of Physiology and Pharmacology, University of Queensland, Brisbane, Australia, and Dr. Barry Gumbiner, Cellular Biochemistry and Biophysics, Memorial Sloan-Kettering Cancer Center, New York, NY) [27]. The negative control used was an equivalent dilution of normal rabbit serum (DAKO). Espin was detected using an immunoaffinity-purified rabbit polyclonal antibody (kindly provided by Prof. J.R. Bartles [24, 28], Department of Cell and Molecular Biology, Northwestern University Medical School, Chicago, IL) with an equivalent dilution of preimmune rabbit immunoaffinity-purified IgG as the negative control. Espin is an actin-bundling protein found only in the ES [24, 28]. Vinculin was also used as a marker for the ES [29] and was detected using a mouse monoclonal antibody raised against human vinculin (clone h-Vin1; Sigma) at a dilution of 1:5000. The negative control used was an equivalent dilution of purified mouse IgG (DAKO).
All incubations were carried out at room temperature in a humidified chamber. All proteins were detected in Bouin-fixed, polyester wax-embedded tissue, except for ß1-integrin, for which frozen sections were used.
The immmunohistochemical protocol has been described in detail previously [24] and used biotinylated secondary antibodies, a streptavidin-horseradish peroxidase complex (ABC complex; Vectastain Elite, Vector Laboratories, Burlingame, CA), and a pink chromogenic substrate (VIP; Vector) to detect the antigen of interest. However, in addition, the signal for PanCad and ß-catenin was amplified by employing a tyramide signal-amplification kit (TSA; NEN Life Science products, Boston, MA). Before chromogenic detection, sections were incubated with TSA-kit streptavidin and horseradish peroxidase solution as instructed by the manufacturer. Sections were then detected with VIP chromogen and processed as described previously [24].
Sections were observed using a 40x objective or a 100x oil-immersion objective on an Olympus BX50 microscope. Images were compiled as described for toluidine blue-stained sections.
The protocol used for double-label immunofluorescence and confocal microscopy has been described in detail previously [24].
Estimation of the Percentage of Tubules with Elongated Spermatids Immunostained for ß1-Integrin, ILK, and Espin during Spermiation
The percentage of seminiferous tubules during spermiation that showed immunostaining for each molecule were determined. Sections were examined at 20x or 40x magnification on an Olympus BX50 microscope, and the image was captured using a Pulinix TMC-6 video camera coupled to a Pentium personal computer using a Screen machine II fast multimedia adapter (Fast, Hamburg, Germany). A software package, DH CASTGRID version 1.10 (Olympus, Munich, Germany), was used to outline the section and to generate a counting frame on the computer monitor. Fields to be assessed were selected by a systematic, uniform, random-sampling scheme with the use of a motorized stage (Multicontrol 2000; ITK, Lahnau, Germany). In each field, the position of a central point in the computer-generated counting frame was recorded to calculate relative stage frequency [19]. Tubules were scored on the presence of adluminal spermatids (i.e., beginning of stage VII to the time of disengagement in mid to late stage VIII) or no adluminal spermatids (i.e. late stage VIII to stage XIV, stages I x VI). Tubules with adluminal spermatids were then further classified as to whether the majority of 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). Approximately 200 points were determined per section. One section per animal was investigated, and four animals per antigen were assessed.
The percentage of tubules with adluminal spermatids was calculated by dividing the number of points landing on tubules with adluminal spermatids by the total number of points landing on all tubules, multiplied by 100. The percentage of tubules with adluminal spermatids that were positive for the antigen of interest were determined by dividing the number of points landing on positive tubules by the number of points landing on all tubules with adluminal spermatids, multiplied by 100.
The difference between the percentage of tubules positive for each antigen of interest was assessed by one-way ANOVA on rank followed by Dunn post-hoc comparison. Data are expressed as the mean ± SD (n = 4 per antigen).
The number of hours between ES removal and spermatid disengagement in control animals (one section/rat, n = 4 rats) was calculated by determining the number of hours in which spermatids associated with the ES during spermiation. The proportion of positive tubules for the ES-marker espin was calculated as described above and then multiplied by 308 h, which is the duration of one spermatogenic cycle [26]. The number of hours in which tubules had adluminal spermatids was calculated by multiplying the proportion of tubules with adluminal spermatids (i.e., stages VII–VIII before disengagement) out of all tubules by 308 h.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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During early spermiation in stage VII, we observed a gradual relocation and concentration of spermatid cytoplasm from around the flagella to form the cytoplasmic lobe that lay near the spermatid head (Fig. 1, A and B). As the cytoplasmic lobe condensed to form a spherical residual body in early stage VIII, the Sertoli cell cytoplasm that surrounds the spermatid head retracted, and the spermatid was extended further into the tubule lumen (Fig. 1C). Disengagement of the spermatid from the residual body and Sertoli cell occurred during mid to late stage VIII. Residual bodies remained after disengagement (Fig. 1D) and were translocated toward the basement membrane.
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In the -T&FSH animals, the process of spermiation appeared to be identical to that seen in control animals. Elongated spermatids were translocated to line the luminal edge in early stage VII, spermatid cytoplasm was removed to form the residual body, and the Sertoli cell appeared to retract from around spermatid heads (not shown). These features were noted with every spermatid examined. The only noticeable difference in -T&FSH testes compared to control testes at the light-microscopic level was that during stage VIII, a significant number of spermatids did not disengage and were retained within the epithelium (Fig. 1E), with both spermatid nuclei and flagella visible within the Sertoli cell cytoplasm. Spermatids that were retained were phagocytosed and translocated to the basement membrane during stage IX (Fig. 1E).
Electron-Microscopic Analysis
Spermatids in early stage VII were associated with ES, which was visible as hexagonally packed actin bundles sandwiched between the Sertoli cell plasma membrane and an underlying layer of endoplasmic reticulum (Fig. 2A). As stage VII progressed, the ES began to be removed, either by breaking down or by removal as a whole unit. Breaking down of the ES was evidenced by partial disappearance of actin bundles and/or endoplasmic reticulum at focal points along the Sertoli cell membrane (Fig. 2A). In some cases, a length of ES could be seen along the Sertoli cell cytoplasm, which had moved away from the spermatid (not shown). Removal of ES in stage VII initially occurred along the ventral side of the spermatid head, because ES was frequently observed along the dorsal side but not on the ventral side (Fig. 2A). The TBCs formed where ES was absent and were characterized by bristle-coated pits invaginating the Sertoli cell cytoplasm (Fig. 2, A and B). The bristle-coated pits extended into the Sertoli cell cytoplasm to form a long, tubular portion (Fig. 2B, inset). The removal of ES and formation of TBCs were seen throughout stage VII and during early stage VIII. In early stage VIII, most spermatids showed little or no associated ES, and the Sertoli cell cytoplasm around the spermatid head had retracted, contacting only small areas of the dorsal aspect of the spermatid head (Fig. 2C). The Sertoli cell cytoplasm retraction occurred predominantly during stage VIII, where little ES was present, and when TBCs were still forming. Disengagement appeared to occur during mid to late stage VIII, and few, if any, spermatids remained at the end of this stage.
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The events described above were seen in each spermatid examined in the -T&FSH animals—that is, ES removal was apparent in all spermatids examined in stage VII, spermatids showed associations with TBCs, and retraction of Sertoli cell cytoplasm was evident from all spermatids observed in late stage VII and early stage VIII (not shown). Spermatids that were retained soon after disengagement in mid to late stage VIII were characterized by an adluminal position and the presence of intact Sertoli cell and spermatid plasma membranes, with no evidence of ES or desmosome-like junctions or of TBC formation (Fig. 2D). Because retained spermatids were phagocytosed by the Sertoli cell and translocated to the basement membrane, Sertoli cell and spermatid plasma membranes disappeared, and the chromatin within the nucleus had a mottled appearance indicative of phagocytosis (Fig. 2E).
The flagella of elongated spermatids during spermiation were also investigated in control and -T&FSH animals. In the -T&FSH testis, there appeared to be no difference in the morphology of flagella in both stages VII and VIII compared to control animals. Similar amounts of Sertoli cell cytoplasm around the flagella in early stage VII were seen (not shown), and spermatid cytoplasm around the flagellum was observed in similar amounts in both control and -T&FSH testes (Fig. 2F).
Immunohistochemical Localization of ß1-Integrin, Cadherins, ß-Catenin, and Espin
ß1-Integrin ß1-Integrin immunostaining was seen around the basal lamina of all tubules at all stages and at sites consistent with the location of Sertoli/Sertoli cell junctional complexes (not shown). Before spermiation, ß1-integrin immunostaining was very faint to absent around elongated spermatids located deep within the epithelium during stages V and VI and at the very beginning of stage VII (not shown). Intense immunostaining of ß1-integrin around the heads of elongated spermatids was seen when spermatids were lined up along the luminal edge some time during stage VII (Fig. 3A). This localization around spermatids persisted into stage VIII (Fig. 3B) and was most notable around the dorsal curvature of the spermatid head (Fig. 3B, inset) in both stages VII and VIII. In late stage VIII, after spermatid disengagement, ß1-integrin immunostaining was not apparent along the luminal edge (not shown). In -T&FSH testes with spermiation failure, the distribution of ß1-integrin was identical to that seen in the controls in all spermatids examined (not shown). ß1-Integrin was also seen around many retained spermatids in stage VIII tubules (Fig. 3C). The localization was most prominent around retained spermatids in the adluminal portion of the epithelium and was less visible as spermatids were phagocytosed and translocated toward the basement membrane.
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Cadherins PanCad immunostaining was evident around the base of the tubule in all stages at sites of Sertoli/Sertoli cell junctions and was associated with developing elongating spermatids before stage VII (not shown). During spermiation, PanCad immunostaining was seen diffusely along the luminal edge associated with elongated spermatids (Fig. 3D). This pattern continued into stage VIII and was associated with elongated spermatids up until disengagement (Fig. 3E). During spermiation failure in the -T&FSH testis, the immunolocalization for PanCad appeared to be the same as that seen in control testes (not shown). Retained spermatids showed faint PanCad immunostaining (Fig. 3F).
N-Cadherin immunostaining at the beginning of spermiation in early stage VII appeared to be confined to elongated spermatid cytoplasm (Fig. 3G) and did not appear to be associated with the Sertoli cell cytoplasm surrounding elongated spermatid heads. As spermiation progressed in stage VIII, N-cadherin localization appeared to be consistent with the position of spermatid cytoplasm and was predominantly seen within cytoplasmic lobes/residual bodies (Fig. 3H), but the localization was not consistent with the presence of N-cadherin in Sertoli cell cytoplasm (Fig. 3H). After disengagement, immunostaining was still seen in the remaining residual bodies (not shown). During spermiation failure (-T&FSH animals), the localization of N-cadherin was consistent with that seen in control animals (not shown). In addition, N-cadherin did not appear to be associated with retained spermatids; however, localization was still seen within residual bodies (Fig. 3I).
ß-Catenin ß-Catenin staining was evident around the base of the tubule in all stages at sites of Sertoli/Sertoli cell junctions (not shown). During spermiation, ß-catenin staining was seen as a diffuse band along the luminal edge in stage VII, which appeared to be confined to the Sertoli cell cytoplasm (Fig. 3J). As spermatid cytoplasm relocated to form the cytoplasmic lobe, ß-catenin appeared most prominently in and around these structures; however, immunostaining was still seen diffusely around elongated spermatids (Fig. 3K). After spermatid disengagement, faint ß-catenin immunostaining was apparent around residual bodies (not shown). During spermiation failure in -T&FSH testes, the localization of ß-catenin was consistent with that observed in control animals at all stages. No ß-catenin immunostaining associated with retained spermatids (Fig. 3L).
Espin Espin, an actin-bundling protein present in the ES, was immunolocalized at sites consistent with Sertoli/Sertoli cell junctional complexes at all stages examined (not shown). During spermiation, espin localization was particularly evident around the head of elongated spermatids during stage VII (Fig. 3M). Around the beginning of stage VIII, espin immunolocalization was no longer associated with the heads of elongated spermatids; rather, staining was prominent in ES structures opposite step 8 round spermatids (Fig. 3N). In -T&FSH animals, the localization of espin was identical to that seen in control animals in stages VII and VIII (not shown). Espin staining was not seen around retained spermatids (Fig. 3O).
Double-Label Immunofluorescence of ILK and Vinculin
Localization of the integrin-associated kinase ILK and of the ES-associated molecule vinculin was examined using dual-label immunofluorescence. During spermiation, the immunolocalization of vinculin was consistent with the position both of the ES and of the staining within the Sertoli cell cytoplasm (Fig. 4A). During stage VII, intense vinculin immunostaining was seen around the entire head of elongated spermatids (Fig. 4A). However, during stage VIII, vinculin appeared to be removed from elongated spermatid heads and was observed mostly in the Sertoli cell cytoplasm, particularly around step 8 round spermatids (Fig. 4B).
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In stage VII tubules, ILK was seen as focal dots in Sertoli cell cytoplasm around the heads of elongated spermatids and appeared to be localized with vinculin close to the heads of elongated spermatids (Fig. 4A). Focal dots of ILK were also apparent along vinculin-labeled strands below the elongated spermatid layer at this stage (Fig. 4A). In stage VIII, ILK was not associated with the elongated spermatid heads (Fig. 4B) but was present in focal dots, along with vinculin, in Sertoli cell cytoplasm around step 8 round spermatids.
The localization of ILK and vinculin and the colocalization of the two proteins were the same in -T&FSH animals (data not shown). Neither ILK nor vinculin associated with retained spermatids during late stage VIII and stage IX (not shown).
Estimation of the Percentage of Tubules with Immunolabeled Spermatids
During stages VII to VIII, ß1-integrin was associated with spermatids in 93% of tubules (Table 1). The remaining 7% that did not show immunostaining associated with spermatids were noted to be tubules in transition between stages VI to VII at the very beginning of spermiation. All stage VIII tubules with adluminal spermatids showed ß1-integrin immunostaining associated with the spermatids. The ILK did not localize with spermatids to the same extent as ß1-integrin, with 67% (P < 0.05 compared to ß1-integrin) of stage VII and stage VIII tubules labeled with ILK. It was also noted that ILK was seen predominantly during stage VII and decreased during stage VIII. Espin staining was seen associated with elongated spermatids in 73% of stage VII and stage VIII tubules. All tubules in stage VII showed espin associating with elongated spermatids, and the 27% of unlabelled tubules were noted to be in stage VIII.
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The time during which tubules had adluminal spermatids was calculated to be 110 ± 10 h (n = 4 animals). The time during which tubules had adluminal spermatids immunolabeled with espin was calculated to be 79 ± 5.9 h (n = 4 animals). Because espin immunolabeling was observed from the beginning of stage VII (see above), it was calculated that ES junctions were removed approximately 30 h before disengagement of the spermatid.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Ultrastructural observations regarding spermiation in control animals in the present study support previous detailed descriptions of this process (for review, see [2]). By examining the series of events in normal rats compared with rats in which suppression of T and FSH causes the failed release of at least 50% of the spermatids [18], the present study suggested that the major defect that occurs during spermiation failure is in the disengagement event. This conclusion is based on the demonstration that in rats with suppressed T and FSH, all spermatids examined during the process of spermiation but before disengagement showed all the normal morphological features associated with spermiation, despite the fact that at least half of these spermatids were destined to be retained by the Sertoli cell.
Previous ultrastructural studies concerning the seminiferous epithelium of hypophysectomized rats suggested that spermatid retention may be associated with the abnormal retraction of Sertoli cell cytoplasm away from the spermatid [16]. Despite detailed analysis, this was not observed by light or electron microscopy in the model of T and FSH suppression used in the present study, and all spermatids examined during early to midstage VIII before disengagement had only a minimal amount of Sertoli cell cytoplasm associated with the dorsal curvature of the spermatid head, as was observed in normal animals. Thus, we propose that retraction of the Sertoli cell from around the spermatid proceeds normally during spermiation failure in our model of T and FSH suppression. In addition, we found no evidence to support the failure of TBC formation during hormone suppression as suggested previously [16]. Although it was not possible to quantify TBC formation, we observed TBCs forming opposite almost every spermatid examined in gonadotropin-suppressed rats. In addition, our observations of spermatids before disengagement and of spermatids retained in the epithelium indicate that removal of spermatid cytoplasm and of the ES junction around spermatids normally occurs during gonadotropin suppression. Because both of these events appear to require TBC formation [2], we believe that failure of TBCs is unlikely to play a major role in spermiation failure. Minor abnormalities observed in the mitochondria of flagella of some spermatids from hypophysectomized rats [16, 30] were also not observed in the present study. The reasons for the differences between our study and the previous studies are unclear, but these differences likely are explained by the fact that hypophysectomy is a more extreme method of gonadotropin suppression and depletes other pituitary hormones as compared to our model.
Morphological observations of elongated spermatids at stage VIII in normal rats of both the present and previous studies [2] show that before disengagement, spermatids do not have ES associated with them. The immunohistochemical localization of the ES-associated molecules espin and vinculin also supports this observation. By determining the proportion of tubules with spermatids immunolabeled for the ES-marker espin, we suggest that the ES is removed approximately 30 h before spermatid disengagement. Therefore, after ES removal, the spermatid is extended well out into the tubule lumen for many hours before release, with Sertoli cell cytoplasm contacting only a small portion of the dorsal curvature of the spermatid head. The components of the intercellular adhesion junction responsible for interaction of the spermatid with the Sertoli cell at this time are unclear. Considering that this junction remaining after ES removal likely mediates disengagement and that disengagement fails during hormone suppression, elucidating the molecular components of this junction is crucial to understanding both normal spermiation and spermiation failure.
Localization of ß1-integrin suggests that it likely is a component of this post-ES junction. ß1-Integrin associated with all spermatids from the beginning of spermiation and persisted until just before disengagement. This localization was restricted to the dorsal curvature of the spermatid only, which is the surface that remains in contact with the Sertoli cell until disengagement. Our findings support the previous demonstration that ß1-integrin is present within the ES junction during spermiation and in developing ES structures opposite step 8 round spermatids [11]. However, our study also suggests that after ES has been removed, as evidenced by negative staining for espin and vinculin, ß1-integrin staining around the spermatid persists, indicating that this adhesion molecule likely mediates spermatid disengagement. This proposition is supported by the demonstration of ß1-integrin immunostaining persisting around retained spermatids after hormone suppression, and it suggests that failure of ß1-integrin-mediated disengagement may be, at least in part, responsible for the failure of spermatid release. Recent studies using an in vitro model of spermiation also provide evidence that ß1-integrin may mediate spermatid disengagement [4]. The authors showed that bacitracin, which inhibits ß1- and ß7-integrin-mediated cell adhesion [31], promoted sperm release from staged seminiferous tubule segments [4]. In addition, concentrations of okadaic acid (a serine-threonine phosphatase inhibitor), which have been shown to increase ß1-integrin phosphorylation and cause it to exit adhesion sites [32], also promoted sperm release in vitro [4].
Previously, ILK has been shown to colocalize with ß1-integrin at sites of ES in the rat testis, and both molecules coimmunoprecipitated from seminiferous tubules [11]. The present study confirms the previous study [11], indicating that ILK interacts with ß1-integrin in ES opposite elongated spermatids in stage VII and in ES developing opposite step 8 round spermatids in stage VIII tubules. Although dual-label immunohistochemistry of ß1-integrin and ILK was not possible because both antibodies were polyclonal, exploring the localization of ILK with the ES-associated molecule vinculin yielded valuable information on ILK in the seminiferous epithelium. The discrete focal dots of ILK seen within vinculin-stained Sertoli cell cytoplasm at sites of ES perhaps suggest that ILK associates with focal aggregates or clusters of ß1-integrin-containing adhesion complexes on the dorsal side of the spermatid. This proposition supports a model recently put forward to explain ß1-integrin and ILK localization in ES structures [11]. In addition, the localization of ILK in filamentous vinculin-stained patterns below spermatids during ES removal in stage VII suggests that these aggregates are removed along with the ES. By stage VIII, when all ES has been removed from around spermatids, ILK and vinculin were not seen around elongated spermatids, despite the fact that ß1-integrin staining persisted. Thus, we believe that ILK likely interacts with ß1-integrin aggregates in association with the elongated spermatid-Sertoli cell ES during spermiation, but that after ES is removed, different ß1-integrin complexes mediate adhesion in an ILK-independent manner until the point of disengagement. The observation of ß1-integrin but not ILK or ES-associated molecules on retained spermatids supports this concept.
In addition to ß1-integrin, cadherins have also been implicated in spermatid adhesion. N-Cadherin is expressed in the testis and has been shown to immunolocalize at Sertoli/Sertoli cell junctions at the base of the epithelium as part of the adherens junction but not the ES [11, 33]. During spermiation, N-cadherin has been previously localized around the curvature of elongated spermatids during stage VII, with this localization redistributing away from elongated spermatids during stage VIII [4, 34], although this was not seen in the present or in previous studies [11, 33], during which localization was not seen around elongated spermatids. N-Cadherin has also been immunolocalized around retained spermatids after boric acid exposure, suggesting that N-cadherin is associated with elongated spermatids during spermiation [4]; however, using the same antibody and similar immunohistochemical procedures, we were unable to see N-cadherin around retained spermatids in our model. Thus, it is unclear if N-cadherin is involved in adhesion during spermiation while the ES is still present, is involved with adhesion of elongated spermatids to Sertoli cells after the ES has been removed, or is involved in disengagement. We suggest that some cadherins are involved in mediating adhesion between the elongated spermatid and the Sertoli cell. A broad-specificity antibody to cadherins (i.e., PanCad) suggested persistence of cadherin until the time of disengagement, with some evidence of immunolabeling of retained spermatids. Localization of the cadherin-associated molecule ß-catenin suggested it was present in a diffuse pattern around elongated spermatids during spermiation, consistent with a Sertoli cell localization. This result is consistent with one other report of ß-catenin localization [12]; however, other studies have shown that ß-catenin is not associated with elongated spermatids [11, 34, 35]. Thus, the role of catenins and cadherins in spermiation remains unresolved.
In conclusion, the present study suggests that spermiation failure after hormone suppression is caused by a dysfunction in the final disengagement of spermatids from the Sertoli cell rather than by a dysfunction in earlier events, such as ES removal, during spermiation. Immunohistochemical data suggest that ß1-integrin, not in association with ILK, plays a role in disengagement by a "loss of adhesion" mechanism. During spermiation failure, this ß1-integrin-mediated loss of adhesion is perturbed, resulting in retained spermatids with associated ß1-integrin immunostaining. Further studies are needed to identify the molecular complex involved in mediating adhesion between spermatids and Sertoli cells after the ES has been removed to identify the cause of spermiation failure induced by hormone suppression.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Liza O'Donnell, Prince Henry's Institute of Medical Research, PO Box 5152,Clayton, Victoria 3168, Australia. FAX: 61 3 9594 6125; liza.odonnell{at}med.monash.edu.au
Received: 6 August 2002.
First decision: 7 September 2002.
Accepted: 23 October 2002.
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REFERENCES |
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