F-actin-based extensions of the head cyst cell adhere to the maturing spermatids to maintain them in a tight bundle and prevent their premature release in Drosophila testis
© Desai et al; licensee BioMed Central Ltd. 2009
Received: 03 April 2009
Accepted: 05 May 2009
Published: 05 May 2009
In Drosophila, all the 64 clonally derived spermatocytes differentiate in syncytium inside two somatic-origin cyst cells. They elongate to form slender spermatids, which are individualized and then released into the seminal vesicle. During individualization, differentiating spermatids are organized in a tight bundle inside the cyst, which is expected to play an important role in sperm selection. However, actual significance of this process and its underlying mechanism are unclear.
We show that dynamic F-actin-based processes extend from the head cyst cell at the start of individualization, filling the interstitial space at the rostral ends of the maturing spermatid bundle. In addition to actin, these structures contained lamin, beta-catenin, dynamin, myosin VI and several other filopodial components. Further, pharmacological and genetic analyses showed that cytoskeletal stability and dynamin function are essential for their maintenance. Disruption of these F-actin based processes was associated with spermatid bundle disassembly and premature sperm release inside the testis.
Altogether, our data suggests that the head cyst cell adheres to the maturing spermatid heads through F-actin-based extensions, thus maintaining them in a tight bundle. This is likely to regulate mature sperm release into the seminal vesicle. Overall, this process bears resemblance to mammalian spermiation.
Spermiogenesis offers a good model for investigating the molecular basis of large-scale cellular morphogenesis and movement. In Drosophila , 64 haploid sperm develop from a single gonial precursor through several well-defined morphogenetic steps [1, 2]. This entire process happens in three distinct developmental stages: (1) the formation of 64 spermatids from a gonial precursor; (2) the elongation of spermatids from nearly spherical to around 1.8 mm long cells; and (3) the individualization of the elongated spermatids into mature sperm, which then enter the seminal vesicle (SV) . Altogether, it involves large-scale changes in cell shape and internal reorganization [3, 4]. Every spermatogonial cell is encapsulated by two somatic-origin cyst cells within the testicular lumen as they form at the testis apex. Subsequent developments occur within this cyst capsule. At the end of the process, the individualized sperm coil up inside the cyst capsule at the base of the testis before entering the SV . Studies in Drosophila indicated that defective sperm fail to enter the SV as they fail to remain attached to the head cyst cell during the coiling process . Thus, the dynamics of sperm head attachment to the cyst cell are likely to play a major role in this quality control exercise.
Spermatids grow in an asymmetric manner and the positions of the nuclei define the rostral ends. Subsequently, the nuclei differentiate into needle-shaped structures containing tightly packed DNA and point towards the SV [1, 4]. As spermatid elongation proceeds, the rostral ends of the nuclei bundle (NB) advance towards the SV at the basal end of the testis. Then, F-actin-based conical structures called investment cones form around each of the needle-shaped nuclei and shortly thereafter, move down the axoneme at a constant speed [4, 5]. This process separately invests each individual spermatid with a plasma membrane, extrudes excess cytoplasm and organelles from the cells, and discharges them as waste bags . Thus individualized, sperm bundles coil up at the base of the testis and then enter the SV [1, 4]. F-actin dynamics, myosin VI, dynamin, dynein light chain 1 (DDLC1/LC8) and myosin V [5–9] as well as several pro-apoptotic proteins [10, 11] are involved in investment cone assembly and the sperm individualization process. The cyst cells also differentiate along with the spermatids. Two cyst cells of different morphological features are found to encapsulate the spermatid bundle at the start of individualization. The head cyst cell caps the rostral ends like a lid on a tube and the rest of the spermatids are enclosed within the tail cyst cell [1, 4]. However, little is known about the molecular basis of sperm release after individualization.
The sperm release process has been extensively studied in the mammalian system. Seminiferous tubules are the functional units of mammalian spermatogenesis. A layer of somatic-origin Sertoli cells line the tubules and constitutes the blood-testis barrier. The Sertoli cells adhere to each other through sets of tight junctions and desmosome-gap junctions [12, 13]. The spermatocytes traverse through these sets of inter-cellular junctions from the basal to the adluminal side as they differentiate (stages 1–7), and complete the differentiation (stages 8–16) while still attached to the adluminal side of the Sertoli cell [13–16]. Then they are released as mature sperm into the lumen. This last stage is called spermiation. Prior to this, each spermatid sheds a residual body containing membrane organelles and cytoskeletal elements, which forms at the junction of the sperm head and flagella . This resembles the shedding of waste bag after individualization in Drosophila testis . Spermatids also associate with the Sertoli cells through cell-adhesion complexes. A testes specific adherens junction (AJ) called an ectoplasmic specialization (ES) is formed at the inner side of the Sertoli cell . This contains characteristic hexagonal actin arrays packed between the ER cisterns and the plasma membrane with a dense formation of tubulin fibers adjacent to the ER [18, 19]. Such a structure has not been reported in invertebrates .
In this paper we report the results of a systematic analysis of the final stages of sperm maturation before their release in Drosophila testis. We show that the somatic-origin head cyst cell grows F-actin based membranous projections into the interstitial spaces between the mature spermatid heads at the start of individualization. Immunohistochemical analysis showed that these F-actin-rich processes contained markers of filopodia and also proteins found in the AJ. Pharmacological manipulations of the F-actin and microtubule dynamics further revealed that these structures are dynamic and are involved in maintaining mature spermatids in a tight parallel bundle. Finally, a genetic screen identified that shibire (dynamin) function is essential to maintain the integrity of these F-actin-based structures and the sperm bundle at the final stage of maturation. Altogether our data provide an initial set of descriptions for further cellular and molecular analysis of spermiation in Drosophila .
F-actin-based membranous extensions of the head cyst cell cover the sperm heads after individualization
Additional F-actin accumulations were observed at the rostral tips of the NBs (arrows, Figure 1B) at the start of individualization. Subsequently, the F-actin grew as cap-like structures around the spermatid nuclei during individualization (Figure 1C). The F-actin densities were found around rostral tips of individual nuclei and acrosomes of an NB (Figure 1D), marked by the sneaky-GFP . At a later stage when the individualized and mature sperm coiled up inside the cyst, the F-actin densities were mostly found around the lateral sides of the nuclei (arrowhead, Figure 1E). This is likely to correspond to a stage when sperm were about to be released as both the acrosomes and the NB appeared unpacked (arrow, Figure 1E). These F-actin based structures will be referred as 'actin caps' in the subsequent discussion. The actin caps were also observed to form inside the head cyst cell covering the rostral ends of maturing spermatid bundles (Figure 1F) and occasionally 'empty' actin caps not associated with the NB were also observed inside the testis. These observations raised an obvious question about the cellular origin of the actin caps.
Cell adhesion proteins along with certain filopodial components are enriched in the actin caps
Filopodia and pseudopodia require the functions of different unconventional myosins [26, 30–33]. We found that both myosin VII (ck) and myosin VI (jar) were present in punctate spots in the head cyst cell cytoplasm (arrows, Figure 4E) and jar/myosin VI was enriched along the actin cap extensions (arrowheads, Figure 4E, b). In addition, expression of the recombinant myosin II regulatory light chain-GFP (sqh-GFP)  also marked the actin caps (arrowheads, Figure 4E, c). Myosin VI is known to stabilize F-actin bundles in the microvilli and the stereocilia , and both myosin II and VIIa were shown to maintain vesicular traffic into the microvilli projections [33, 35]. In Drosophila ovary, myosin VI was localized in the membrane protrusions at the leading edge of migrating border cells , and at the investment cones in the testis . Therefore, the actin caps appeared to be filopodia-like projections of the head cyst cells maintained by myosin-dependent vesicular traffic. However, we noted that unlike the ck:GFP, the ck antibody did not label the actin cap extensions. The ck antibody is raised against the stalk domain of the predicted myosin VII ORF , which is likely to be involved in homodimerization of the motor subunits and also in binding other accessory subunits. Therefore, unlike the ck:GFP, the antibody could only label myosin VII (ck) in the tissue if the epitope is exposed. Furthermore, the ck:GFP transgene is ectopically expressed using a non-homologues promoter and this could also alter its subcellular localization in the cell. All of these could account for the apparent differences in the subcellular staining patterns of ck in the head cyst cell.
A large number of vesicles were reported to accumulate inside the head cyst cell around the rostral tips of the embedded sperm heads . This was also revealed in the DIC images of the isolated head cyst cells. The punctate distributions of myosins in this region further suggested that these motors might be involved in vesicle transport into the actin cap projections. Myosin VI was known to associate with the clathrin-coated pits inside the cell during endocytosis as well as with dynamic membrane ruffles in the migrating epithelial cells . We found that the clathrin light chain-GFP (clc-GFP) was enriched in punctate spots around the actin caps (arrows, Figure 4E, d) in the w; UAS-clc-GFP Actin5cGal4 testis. Clathrin is involved in coated vesicle assembly  and plays an important role in dynamin-mediated vesiculation inside the cell and the clc is an integral part of the clathrin complex. Therefore, this observation further supported the hypothesis that vesicular traffic from the head cyst cells could supply membrane to the actin cap projections. Altogether the immunolocalization results indicated that actin caps are filopodia-like extensions and likely to attach to the sperm heads during individualization.
F-actin and microtubule stability are essential for actin cap assembly and maintenance
Relative NB and IC disruptions in individual testis after 30 minutes of lat B (25 μM) treatment.
NB disruption (%)
IC disruption (%)
shibire/dynamin is essential for maintaining the actin caps and spermatid bundles
The association between head cyst cells and spermatids was predicted to play an important role in spermiation . The above results indicated that the head cyst cells are likely to adhere to the maturing sperm heads through F-actin-rich projections containing several filopodial markers. Dynamin plays a key role in membrane reorganization process and our immunolocalization data suggested that it could play an important role in actin cap assembly or maintenance as well. The shibire gene codes for a dynamin homologue in Drosophila , which is shown to play an important role in endocytosis  and various other cellular functions . Previous studies have also shown that shi ts1 (a conditional mutant allele of shibire ) hemizygous flies rapidly paralyze within a few minutes at non-permissive temperatures . Thus, the conditional shi ts1 alleles provided a good tool to further test the role of dynamin in actin caps and the latter's role in sperm maturation/release.
shibire/dynamin function is required in the head cyst cells to maintain the actin caps and the sperm head bundles
Actin caps organize spermatid heads in a tight bundle inside the head cyst cell
Post individualization, all of the 64 clonally derived spermatids are twisted together and the bundle coils up inside the cyst. The head cyst cell attaches to a terminal epithelial cell at the base of the testis, and then the sperm is released into the SV . Our studies identified a specific function of the head cyst cell in the sperm bundling process during the maturation stage. As summarized in Figure 9B, it is found to grow several F-actin-based extensions at the start of individualization that are enriched with proteins found in the filopodia of other cell types . In addition to dynamin, syndapin and WASP, proteins found in the apical ES [54, 55] such as ERp72, E-cadherin and beta-catenin are enriched in the actin cap extensions. This suggests that the actin cap extensions are dynamic membrane bound projections and they adhere to the individualized sperm heads while the remaining parts of the spermatids undergo morphogenetic changes during the individualization and coiling stages. Lamin, a protein associated with the F-actin cones during the sperm individualization process  was enriched at these structures. This could strengthen the membrane cytoskeleton and provide rigidity to the actin cap extensions. Myosins are essential for the formation of filopodia-like extensions  and for cell adhesion [56, 57] in several other cell types. Hence, the presence of the myosin II regulatory light chain, Myosin VI and myosin VII suggests frequent vesicular membrane transport into and out of the actin cap projections as well as cross linking of actin filaments to the plasma membrane inside these extensions. The presence of clathrin light chain and syntaxin further pointed towards the existence of dynamic membrane exchange along the actin cap projections. Altogether these immunolocalization studies suggested that actin cap extensions could adhere to the maturing sperm heads during individualization and coiling through a combination of different cellular mechanisms. This is also confirmed by the results of our pharmacological and genetic analyses, which indicates that the actin cap extensions and their interactions with the spermatid heads are dynamic. Altogether, it showed that the differentiating spermatids are tightly attached to the head cyst cell until they are fully mature and suggested that this could play a critical role in preventing premature sperm release inside the testis.
F-actin-based membrane remodeling at filopodia and pseudopodia-like structures is involved in cell migration, axonal growth cone guidance and cell ingestion . In Drosophila ovary, the somatic origin 'border cells' migrate through the germ-line 'Nurse cells' by extending similar F-actin-based membranous projections . Actin-based cellular extensions were also known to anchor epithelial cells on the substratum involving beta-catenin and cadherin . White blood cells migrate out of the blood vessel by penetrating the endothelial cell layer through such actin-based membranous projections by a process called transcytosis . Drosophila spermatids lose all of their cytoplasmic contents after individualization by the movement of the F-actin-based investment cones through them [3, 7]. Hence, the somatic-origin cyst cells that encapsulate these near-mature sperm are expected to play an important role in regulating their release. This is further established by a targeted disruption of shibire/dynamin functions in the head cyst cells, which indicate that the maintenance of spermatids in a tight bundle is essential for preventing a premature release. This is also consistent with observations made in the mammalian testis. The Sertoli cells play an important role in the mechanical movement of germ cells from the basal to the adluminal side during their differentiation . This process is aided by the F-actin-rich ES and a variety of different cellular junctions [14–16]. Our results helped to identify the molecular and cellular framework involved in the spermiation process in Drosophila . This also provides a useful base for future analysis of sperm release mechanisms.
Dynamin-mediated membrane remodeling is essential for actin cap maintenance and attachment to sperm nuclei at the terminal stages of maturation
Dynamin is implicated in the assembly and maintenance of F-actin-based membrane ruffles, podosomes and invadopodia [60–63], and a specific dynamin isoform in rat (dyn3) was found to associate with the tubulobulbar junctions around the sperm heads [64, 65]. Dynamin is enriched at the actin caps and genetic analysis showed that they are essential for maintaining the sperm bundle inside the cysts. The dynamin requirement appeared to be constitutive as the temporal loss of shibire/dynamin function caused actin cap disassembly and NB disorganization. In addition, it caused accumulation of NBs inside the testes within 30 minutes. Thus, dynamin-mediated membrane remodeling at the actin cap region is expected to play a role in sperm bundling and release processes in Drosophila , and provided a basis to investigate the interactions between the somatic origin cyst cells and spermatids during the sperm maturation process in Drosophila.
In view of the observations presented here, the sperm maturation process in Drosophila resembled the spermatid development inside the Sertoli cell layer in mammalian testis . The mammalian spermatids attach to the Sertoli cell membrane through the apical ES . The F-actin-based ES forms around developing spermatids inside the Sertoli cell . Although electron microscopic studies did not reveal ES-like structures around the sperm heads in Drosophila testis, the actin caps contained the essential functional elements of these specialized junctions. It is rich in F-actin, beta-catenin and DE-cadherin, and tightly associated with the spermatid heads during individualization. In addition, we found a second interesting parallel. Mammalian sperm are physically released from the Sertoli cells after the removal of integrin and beta-catenin from the apical ES . A tubulobulbar junction forms inside the Sertoli cells and around the mammalian spermatids [54, 55]. Proteins involved in endocytosis, such as dynamin 3 and amphiphysin were enriched at these junctions [64, 65, 68], and loss of amphiphysin from the tubulobulbar junctions was shown to block sperm release in the knockout mice . We showed that shibire/dynamin functions in the head cyst cells in Drosophila testis to maintain the actin cap integrity and sperm heads in a tight bundle. Thus, our observations have the potential to establish Drosophila as an attractive model for molecular analysis of spermiation in insects. It has defined an assay to study the role of F-actin-mediated cell adhesion process and can be used to screen for small molecule-based perturbation of the sperm maturation process in the future.
Drosophila stocks and culture conditions
All of the fly stocks used for this study are listed in Additional file 6. They were maintained on standard cornmeal agar medium as described previously . We sincerely acknowledge the generous gifts of fly stocks from the respective sources. For most of the immunostaining and analysis, 2-day old adult males grown at room temperature were used. The conditional mutant alleles of shibire/dynamin (shi ts1 and shi ts2 ) were grown at 18°C and 2-day old adults were shifted to either 29°C or 32°C for a defined period before dissection and immunostaining. All of the transgenic fly stocks were grown at 25°C. The crosses set for the heat pulse studies were grown at 18°C. Subsequently, the freshly emerged males were maintained at the same temperature for 2 days before they were shifted to the non-permissive temperatures as per the requirements.
For whole mount analysis, testes were dissected in phosphate buffered saline (PBS)  containing 0.01% saponin (Sigma Chemical Co., MO, USA), incubated in the same solution for 30 minutes and then fixed in PBS containing 4% paraformaldehyde (freshly prepared). After several quick rinses in PTX (PBS with 0.3% Triton X-100), the fixed testes were incubated in different primary antibody solutions in PTX for 1 hour at room temperature. This was followed by washes in PTX, and further incubation in appropriate fluorescent labeled secondary antisera in PTX for 1 hour. After a final series of washes, the tissue preparations were mounted on a glass slide with a drop of Vectashield® (Vector Laboratories Inc., USA) and under a #1 cover glass. To observe the nuclei and the F-actin distribution in the tissue, the immunostained specimen were additionally incubated in PTX containing 76 μM of FITC:phalloidin or RITC:phalloidin (Sigma Chemical Co., MO, USA) and 0.001% DAPI (Sigma Chemical Co., MO, USA) for 30 minutes, washed in PTX and then mounted as above. In some cases individual cysts were teased out of the testis after staining and dispersed on the slide before mounting. This helped to reveal the subcellular distribution of F-actin structures. For squash preparations, the testes were dissected on a plus charged slide, squashed under a cover slip, dipped in liquid nitrogen for a few minutes and then in ice-cold 95% ethanol for 3 minutes, or, until the cover slip dropped off. This was followed by a post fixation of the slide in PBS containing 4% paraformaldehyde and immunostaining as per the procedure described earlier . This often disrupted the head cyst cells but kept the NBs intact. In all of these cases the actin caps were found to remain associated with the intact NBs.
For the pharmacological treatments, adult testes were dissected in PBS containing 0.003% DMSO and either 30 μM lat B or 5 μM vinb (both from Sigma Chemical Co., MO, USA), and then incubated in the same media for 30 minutes before fixation. Then they were processed as described above. For short heat pulses, dissected testes in PBS were incubated in a water bath set at defined temperatures (18°C for controls and 29°C for heat pulse) for the specified period of time. Then they were processed as described above.
A list of all of the antibodies used for this study and their respective sources is provided in Additional file 7. All of the fluorescently conjugated secondary antibodies were obtained from the Jackson Research Laboratories Inc., USA, and from the Molecular Probes Inc., OR, USA, and used at 1:400 dilutions.
Testes were dissected from 2-day old males in 2.5% glutaraldehyde (EM Sciences Inc., USA), 4% paraformaldehyde and 0.04% CaCl2 in 0.1 M phosphate buffer (pH 7.4) at 4°C, then fixed overnight in the same solution, washed in 0.1 M phosphate buffer (pH 7.4) and post-fixed in OsO4 for 4 hours at 4°C. This was followed by several washes in 0.1 M phosphate buffer (pH 7.4), dehydration in graded series of ethanol, and embedding in Araldite (E. Merk GmbH, Germany). Ultrathin (100 nm) sections were obtained in Leica Ultracut 6b, stained with aqueous uranyl acetate and lead citrate, and imaged in a JEOL 100S electron microscope as per the procedure described previously .
Statistical analyses of the NB and IC morphologies in the wild type and mutants
The number of mature NBs and the ICs were counted in each testis preparation under a 40 × 0.75 NA objective fitted in an epifluorescence microscope. The NBs and the ICs were carefully scrutinized for abnormal organizations and if they were found to be out of register with each other, then they were counted as disrupted. The principal criteria used for this analysis has been described earlier . Briefly, the criterion for a NB to be considered as intact, most of the nuclei should remain in parallel register and be packed tightly together (example: Figure 6C, a). Only those which appeared obviously disrupted were counted as not intact (example: Figure 6C, b and 6C, c). The F-actin cones in an IC are found in a parallel register (example: Figure 6E, a) while the disrupted ones had them scattered to different extents (example: Figure 6E, b and 6E, c). Volunteers also counted some of the preps, selected at random, in a double blind manner. This showed that the criteria used for counting were quite robust. The results were presented as histogram plots with ± standard error of the mean, and the significance of the differences was calculated by using the Mann-Whitney non-parametric test using the Graphpad Instat™ software.
The NB and IC defects as well as the total numbers were recorded from 10 lat B treated testes. The relative NB and IC disruption values from individual testis were plotted on the x - and y -axis, respectively. The data was then analyzed by using Graphpad Instat™ and the SigmaPlot™. Graphpad Instat™ was used to determine the pairwise significance (p ) values and to estimate whether the slope was significantly different from zero. SigmaPlot™ was used to determine the slope of the best-fitted line amongst the points in the plot. For studying the correlation between different genotypes, the average NB and IC disruption values (determined as a percentage of the total) from each genotype were plotted and analyzed as described above.
Image collection and analysis
All images were collected by using the Olympus FV1000SPD laser scanning confocal microscope (LSCM). The image frames were merged by using ImageJ® http://rsb.info.nih.gov/ij, and adjusted for their brightness and contrast to maintain uniform visibility in a montage by using Adobe Photoshop® (Adobe Corp., USA). The figures were then organized and labeled in Adobe Illustrator™.
clathrin light chain
differential interference contrast
green fluorescent protein
- lat B:
monomeric red fluorescent protein
nuclei bundle of maturing spermatids
transmission electron microscopy
upstream activating sequence
Wiskott-Aldrich syndrome protein.
We thank Barbara Wakimoto, Dan Kiehart, John Roote, and Eyal Schejter for generous and prompt sharing of fly stocks and reagents. We thank Vimlesh Kumar and Mani Ramaswamy for providing the syndapin antibody. P. Koshire helped KR in setting up genetic crosses and in sample preparations. BSD and KR also acknowledge the summer students, D. Kar, A. Sarkar and K. Mukherjee, who helped in the initial stages of this project. BSD thanks A. Ghosh Roy for valuable discussions. Research described in this article was supported by an intramural grant from TIFR, India to KR and a TIFR scholarship to BSD.
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