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Vol. 19, Issue 6, 2363-2372, June 2008
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Department of Biology, Washington University in St. Louis, St. Louis, MO 63130; Faculty of Biology and Earth Sciences, Institute of General and Molecular Biology, Laboratory of Developmental Biology, Nicolaus Copernicus University, 87-100 Torun, Poland; and *Laboratory for Morphogenetic Signaling, Center for Developmental Biology, RIKEN Kobe, Kobe 650-0047, Japan
Submitted August 27, 2007;
Revised February 20, 2008;
Accepted March 4, 2008
Monitoring Editor: David Drubin
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Ridley et al., 2003; Carlier and Pantaloni 2007). Several different types of actin structures are known to contribute to motility. Filopodia, which contain parallel bundles of actin filaments, are thought to be important for exploring the cells' environment and making initial contacts with substrate for movement in a particular direction. Lamellipodial meshwork is proposed to push the leading edge of the cell forward using the force generated by addition of actin monomers at the barbed ends of the filaments, which face the membrane (Pollard and Borisy 2003). Behind the leading edge, a lamellar region consisting primarily of bundles that contain tropomyosin and myosin II plays an important role in cell movement (Gupton et al., 2005). In addition, stress fibers, composed of antiparallel actin filament bundles connected to adhesion complexes, are important for traction and contractile forces that release the cell–substrate attachments so that forward movement can occur.
The different actin structures and domains are regulated by different sets of actin-associated proteins. Much is known about control of assembly of the two main types of actin organizations, meshwork and bundles, from biochemical analysis in vitro and studies of motile cells in vivo. Actin meshwork is nucleated by the Arp2/3 complex, and the branched organization relies on the ability of the complex to bind to the side of an actin filament and nucleate a new filament (Goley and Welch, 2006). Parallel bundles are nucleated by formins, which associate with the barbed ends, but allow monomer addition while remaining bound (Goode and Eck, 2007). For some formins, barbed-end monomer addition requires the presence of profilin (Romero et al., 2004; Kovar et al., 2006), a monomer binding protein. In the case of both meshwork and bundles, other proteins work in conjunction with the nucleators to further regulate organization, polymerization dynamics, attachments, and other aspects important for function.
The mechanisms that control the formation and maintenance of actin structures in specialized cell types are less well understood. Actin structures in differentiating and differentiated cells are important for shape, connections to other cells and the extracellular matrix, and the cell's specialized features that are involved in its physiological roles in the context of the tissue and organism (Revenu et al., 2004). In general, actin structures in differentiated cells appear to be organized based on the same principles as the lamellipodial meshwork, filopodial and lamellar bundles, and stress fibers, but often they are significantly different in size and dynamic properties than those in motile cells (for some examples see Tyska and Mooseker 2002; Tilney and DeRosier 2005; Sekerkova et al., 2006). To understand how the principles and activities that have been characterized using motile cells as models can be applied widely to actin in the context of differentiated cell function, exploring the formation and function of a variety of the specialized actin structures found in a number of different cell types is important.
One model system that presents an interesting case of intracellular motility in the context of a developmental process is Drosophila spermatid individualization. Preceding individualization, the germline precursors of the sperm undergo mitotic and meiotic divisions with incomplete cytokinesis to generate cysts of 64 syncytial spermatids. The cysts elongate and elaborate axonemes. Finally, the syncytial spermatids are separated into individual cells in a process called individualization. Individualization reorganizes the syncytial spermatid membrane and removes most of the cytoplasmic contents (Tokuyasu et al., 1972; Fabrizio et al., 1998; Hicks et al., 1999; Noguchi and Miller 2003). Structures called actin cones mediate this cellular remodeling. Actin cones assemble around the sperm nuclei and travel away from the nuclei along the length of the axoneme, synchronously at constant speed, over a distance of 
2 mm, taking 10 h. The pushing out of the cytoplasm and organelles by the actin cones during individualization results in the formation of a "cystic bulge" of accumulated cell contents ahead of the cones. After individualization is complete, the membrane completely and tightly encloses each nucleus, axoneme, and mitochondrial derivative complex, and most of the cytoplasm and organelles are gone.
We have previously studied the individualization process (Hicks et al., 1999; Rogat and Miller, 2002; Noguchi and Miller, 2003) and determined that the actin cones have two domains, a rear region of parallel bundles and a front region of meshwork (Noguchi et al., 2006). We also showed that myosin VI is important for stabilization of the actin cones as they move (Noguchi et al., 2006). In the present work, we examine how and when each part of the cone is formed and the function of each part during cone movement and cellular reorganization. We find that the two regions form by different polymerization mechanisms, at different times, and play different roles in cone movement and individualization. Surprisingly, the meshwork is not required for movement. Instead, the bundles are important. By analogy to a steam locomotive, the bundles serve as the locomotive and the meshwork serves a structural role, functioning similarly to a "cow catcher," pushing the cytoplasm and organelles in front of it. This system provides an example of how the basic mechanisms in play in motile cells are modified to achieve a different result in the context of cellular differentiation in vivo.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Antibodies
Anti-D. melanogaster-arp3 antibody was a kind gift of Bill Theurkauf (University of Massachusetts Medical Center, Worcester, MA; Stevenson et al., 2002). Anti-quail and anti-singed antibodies were made by Lynn Cooley (Yale University; New Haven, CT) and obtained from the Developmental Studies Hybridoma Bank. Anti-myosin VI mAb (3C7) was described previously (Kellerman and Miller, 1992).
Isolation and Primary Culture of Cysts
Primary culture of individualizing cysts was carried out as previously described (Cross and Shellenbarger, 1979; Noguchi and Miller, 2003). The arp3 mutation causes lethality at a late pupal stage (Hudson and Cooley, 2002), so testes could not be obtained from adults. Therefore, live homozygous arp3 mutant pupae were identified by the lack of the Tb (short body length) marker, which is present on the balancer chromosome, at a late stage (with dark visible wings) when spermatogenic cysts were viable in culture. The pupal arp3 mutant testes were dissected and contained some elongated cysts and some individualizing cysts. Elongated cysts were dissected and cultured.
Immunofluorescence Microscopy
Immunostaining of isolated spermatogenic cysts from various mutants was performed as described previously (Noguchi and Miller, 2003). Specimens were examined using laser scanning microscopy (LSM; Leica, Iyna, Germany) with a 40x lens at 4x digital zoom. For DNA/F-actin staining, DNA was stained with 1 µM DAPI and F-actin was stained using Alexa-568-phalloidin. Specimens were examined with a Nikon inverted microscope equipped with a cooled CCD camera (CoolSNAP ES, Photometrics, Woburn, MA) driven by Metavue software (Universal Imaging, West Chester, PA).
Quantitation of F-actin in actin cones before the onset of individualization was performed by measuring fluorescence intensity of actin cones in isolated cysts. Actin cone bundles associated with sperm nuclei as judged by F-actin/DNA staining were selected for examination. Images were obtained using an Olympus ASW LSM (Melville, NY) with a 10x lens, and the signal intensity of actin cones was measured and processed using ImageJ (http://rsb.info.nih.gov/ij/) and Excel software (Microsoft, Redmond, WA). Quantitation of actin amount in cones that had moved was performed as previously described (Noguchi et al., 2006). Length and width of actin cones that had moved was measured in high-magnification images (40x lens) using ImageJ software. Cones that were not associated with nuclei were selected for examination. Student's t tests were performed to evaluate the significance of differences in measurements between genotypes.
Electron Microscopy
For cross sections of spermatogenic cysts, dissected testes from adult male flies were fixed with 1.5% glutaraldehyde, postfixed in 1% OsO4, and embedded in PolyBed 812 resin (Polysciences, Warrington, PA) using the procedure described previously (Noguchi et al., 2006). Stained sections (60–70 nm) were cut using a Leica UTC ultramicrotome and then examined using a JEOL EM 1010 transmission electron microscope (Peabody, MA).
Myosin Subfragment 1 (S1) Fragment Decoration
Purification of rabbit skeletal myosin II and preparation of S1 subfragment were carried out using conventional methods (Margossian and Lowey, 1982). For myosin II S1 fragment decoration, isolated cysts were permeabilized with 0.1% saponin and treated with 4 mg/ml S1 fragment using the procedure described previously (Noguchi et al., 2006). For this study we used cysts that were classified into three different stages by examination of morphology under a dissection microscope: 1) before cone movement, a very early stage without any signs of cystic bulge formation, 2) an early stage of individualization, with a small cystic bulge near the end of the cyst, and 3) individualizing cysts with the cystic bulge positioned between one-fourth and one-third of the cyst length. To obtain the five samples for which actin cone organization in early cones could be seen that were used in this study, we performed six to seven experiments. In each experiment testes were dissected from 50 males, and 60–70 cysts at the right stage were selected. Although the cysts were handled carefully under a dissection microscope, the majority of them were lost during the long procedure. In addition, only a subset had actin cones. In most cases one preparation yielded one to three S1 decorated samples at the end. After sectioning, some samples were not oriented properly to see cones in sections, resulting in only a few cysts that yielded results.
chic7886/chic7886 mutant cysts do not form any cystic bulges, so individualizing cysts cannot be identified. To identify cysts that had cones, actin was stained with Alexa488-phalloidin during extraction with 0.1% saponin. Cysts with actin staining were selected using a fluorescence dissection microscope, and S1 decoration was performed as previously described (Noguchi et al., 2006). S1-decorated cysts were fixed with 1% glutaraldehyde and 0.2% tannic acid, postfixed in 2% OsO4, and embedded in Poly/Bed 812 resin as described previously (Noguchi et al., 2006). Ultrathin sections were cut, stained, and examined as described above.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Such cysts were fixed and processed for myosin S1 fragment decoration and EM visualization as previously described (Noguchi et al., 2006). Cysts were positioned so that the ends where the nuclei were present were sectioned longitudinally. We saw two different organizations of actin in cones of early cysts. In most early cysts, the cones contained only bundles (four of five early cysts, 12 cones). In these cones, loose bundles of actin filaments surrounded the nuclei (Figure 1, A–C). On the side of the nuclei away from the direction of cone movement (large arrow in Figure 1A), the space between the syncytial membrane (arrows, B and C) and the nuclei was small, and the whole area was filled with actin bundles that ran parallel to the long axis of the cyst (Figure 1, B–D). Along the sides of the nuclei, the membrane was also close and the area was filled with parallel bundles. In the region in front of and further from the nuclei, where axonemes/mitochondria were observed, there was a slight shift in the actin (Figure 1C, front). Some of the filaments in this region did not lie parallel to the long axis of the cysts, but instead were at an angle (Figure 1, C and E).
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We evaluated filament orientation in both of these types of cones. We quantitated filament orientation by classifying filaments in the front and rear domains as parallel or perpendicular to the long axis of the cyst, and noted the direction of the barbed end, as determined by S1 decoration (Figure 2A). We determined that the filaments were primarily oriented with barbed ends facing away from the direction of movement and toward the membrane surrounding the back of the cone in both cones with no meshwork and cones with a small amount of meshwork (Figure 2B). This is similar to our observations of filament orientation in moving cones (Noguchi et al., 2006). These data show that overall orientation of the filaments is the same throughout assembly and movement.
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). Thus, before movement began, the cones had no meshwork and were entirely made up of bundles. The meshwork formed coincident with initiation of movement.
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). If the Arp2/3 complex is important for meshwork formation, we would predict that it should become enriched at the cone front, where the meshwork is formed, upon initiation of movement. We would also predict myosin VI would become enriched at the front coincident with Arp2/3 complex, because myosin VI function is required for Arp2/3 complex accumulation there (Noguchi et al., 2006). We quantitated the distribution of arp3 (representing Arp2/3 complex localization) and myosin VI on cones that had not yet moved and cones that had initiated movement. Cones that had not moved were identified as those with a rectangular shape that were associated with DAPI-stained nuclei. Myosin VI (Figure 4Aa) and arp3 (Figure 4Ab) were present at levels above background in about half of the population of cysts that contained cones with these characteristics (myosin VI: 47 ± 9%, arp3: 54 ± 14%; n = 3 experiments; 30 cysts in each experiment). Among these, the vast majority showed uniform staining along the length, with no concentration on the front (myosin VI: 88 ± 6%, arp3: 100%).
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30 cysts in each experiment) and arp3 (Figure 4Bb; 88 ± 17%) at the front. Only a very few showed no concentration at the front with staining evenly spread over the cones (not shown; myosin VI: 3 ± 3%; arp3: 4 ± 4%). The rest showed no concentration above background. The polarized distribution was maintained throughout movement. This tight coupling of polarization of these actin regulators with the onset of movement is consistent with our observations above, that formation of the meshwork, which is likely to be an Arp2/3 complex–nucleated process, shows strict correlation with the start of movement.
Because the rear of the cones is composed of actin bundles, it seemed likely that actin-bundling proteins would be present in this domain. We examined the distribution of two actin-bundling proteins: quail (Mahajan-Miklos and Cooley, 1994) and singed (Cant et al., 1994). Quail is an ortholog of villin that bundles actin filaments, but does not sever in vitro and is important for bundle formation during oogenesis (Mahajan-Miklos and Cooley, 1994), whereas singed is a fascin ortholog that is important for actin bundle formation both during oogenesis and in bristles (Cant et al., 1994). As cones formed and before movement, when the cones contained only bundles, both quail (villin) and singed (fascin) were present all over the cones (Figure 4A, c and d). Quail (villin) is concentrated at levels well above background on the cones at this early stage, whereas singed (fascin) appears to be less enriched. After movement began, both these proteins' distributions changed (Figure 4B, c and d). Both were enriched in the rear region where bundles were present and absent from the front region where the meshwork was present. However, their distributions were not identical. Quail (villin) occupied the middle portion and singed (fascin) localized to the extreme rear of the cones (compare the overlays in Figure 4B, c'' and d''). It is likely that these actin-bundling proteins contribute to formation and stabilization of actin cone bundles. The distributions of these four actin-associated proteins (quail, singed, arp3, and myosin VI) show that the two structural domains have functionally different actin-regulating proteins associated with them. The proteins present in each domain have activities that are consistent with a role in generating and/or stabilizing those domains. These results also reveal that the actin cone structure is complex with at least four distinct regions based on protein composition (see schematic diagrams, Figure 4, C and D).
The Front Meshwork Requires Arp2/3 Complex to Form
Because of the very different organization at the front and rear, we anticipated that they would be built using two different actin polymerization mechanisms. The front meshwork structure is consistent with Arp2/3 complex–mediated branched network formation. We disrupted Arp2/3 complex function using a mutation in the arp3 gene. The arp3 mutation is homozygous lethal at the pupal stage, so we could not examine testes from adults. However, if larvae are grown in optimal conditions, the homozygous mutant animals survive until the very late pupal stage. Late stage pupae were dissected to obtain testes, and cysts were isolated and cultured. Fully elongated cysts were present and individualizing cysts could be observed, but individualization was abnormal (see Supplemental Data, movie of individualization of arp3 mutant cysts, and below). F-actin staining revealed that arp3 mutant cones in individualizing cysts were not normal in shape, but instead appeared very narrow, much more like cones in wild type that had not initiated movement (Figure 5Aa'). Measurement of the width of moving cones in the arp3 mutant supported the idea that they are narrower than normal (arp3 mutant: 1.0 ± 0.3 µm [n = 12]; wild type: 1.8 µm ± 0.3 [n = 23]; p < 0.0001). arp3 mutant moving cones also contained less actin than wild-type cones, as shown by the intensity of actin staining (relative amount of actin staining: arp3/wt = 0.60 ± 0.3; n = 15 cones; p < 0.005). However, before the initiation of movement, arp3 mutant cones had an equivalent amount of actin as wild-type cones (relative intensity of actin in arp3/wt = 0.92 ± 0.36, n = 11; p > 0.5). The fact that the cones form normally before movement initiates when only bundles are present, but have less actin after movement begins when the meshwork normally grows, is similar to myosin VI mutant cones (Noguchi et al., 2006) in which the meshwork does not grow properly. Immunofluorescence localization of actin regulators showed that these distributions were altered in the arp3 mutant. Myosin VI was not concentrated on the cones or localized at the front (Figure 5Aa), quail (villin) was present from the front to approximately the middle (Figure 5Ab), and singed (fascin) occupied the rear half of the cone (Figure 5Ac). Because bundling proteins were observed along the entire length of the cone and cone shape was not triangular, we conclude that the front meshwork was greatly reduced or absent in arp3 mutant cones. Direct observations of filament organization using myosin II S1-decoration at the EM level were not possible in the arp3 mutants, because we could not reliably identify cystic bulges to find moving cones (see the Supplemental Movie for an example). In addition, it was difficult to collect a large number of individualizing cysts from the arp3 mutant, due to the lethality of this genotype.
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), so testes could be isolated from the mutant. This mutant has cytokinesis defects during earlier spermatocyte divisions (Giansanti et al., 1998), but these defects do not prevent elongation. Numerous fully elongated cysts with mature sperm nuclei were observed at the basal end of the testis, but almost no actin cones were detected along the length of the testis where they normally can be seen as they move along the axoneme during individualization (see below). chic (profilin) mutant cones were almost always associated with nuclei at the basal end of the cyst. chic (profilin) mutant cones had a very different appearance (Figures 5B and 7A) from wild-type cones. In cysts that correspond to the stage when the cones initially form, chic (profilin) mutant cones were significantly smaller than wild type (see Figure 7A). Quantitative analysis of F-actin in cones at this stage demonstrated that the amount of F-actin is reduced to one-third of wild type (0.32 ± 0.06, p < 0.0001; n = 3 experiments, 10 cysts in each experiment). Because during early stages of cone formation, the cones consist of only actin bundles, the small amount of actin present is consistent with the idea that profilin is important for polymerization of the bundles. Some cones associated with nuclei had a short triangular shape with a very wide front (Figures 5B, d'–f', and 7B). The length of these chic (profilin) mutant actin cones was significantly shorter than wild-type moving cones (actin cone length: wild type, 12.0 ± 1.21 µm, n = 23; chic7886/chic7886, 6.1 ± 1.6 µm, n = 23, p < 0.0001). However, the width was comparable (width: wild type, 1.8 ± 0.3 µm; chic7886/chic7886, 2.0 ± 0.4 µm, p > 0.05). Myosin VI (Figure 5Bd) and arp3 (Figure 5Be) localized normally relative to each other on these triangular-shaped cones. However, arp3 occupied the whole length of the cones (Figure 5B, e–e''). In addition, singed (fascin; not shown) and quail (villin; Figure 5Bf) were barely detectable on the mutant cones. The lack of bundling proteins, the short length, and the distribution of arp3 along the entire length of the cone suggest that the rear region of bundles was greatly reduced in chic (profilin) mutant cones. S1 decoration demonstrated that chic (profilin) mutant cones that were associated with nuclei contained a large amount of meshwork and almost no bundles (Figure 6). In wild-type cysts, cones associated with nuclei were composed of a large number of bundled filaments and rarely had any meshwork at all (see Figure 1). We never observed any cones associated with nuclei with extensive meshwork in wild type, as seen in the chic (profilin) mutant.
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In cultures of cysts from chic (profilin) mutants, we did not observe any cystic bulge formation or signs of individualization. In addition, we rarely observed cones that had moved away from nuclei, and cones associated with nuclei were often triangular in shape (Figures 5B and 7B), like moving cones in wild type. Because we showed above that profilin mutant cones have a greatly reduced amount of bundles and contain mostly meshwork, we conclude that cones with a deficit of bundles cannot move.
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and T. Noguchi, unpublished observations).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To our knowledge, this Arp2/3 complex–nucleated meshwork's role in mediating cytoplasm exclusion is novel. This meshwork is dense enough to completely exclude cytoplasm from a compartment of the cell. To form such a dense structure, myosin VI stabilization could be important. Myosin VI mutant cones are not as dense as normal and do not effectively exclude cytoplasm and organelles (Noguchi et al., 2006). Very dense actin meshwork might be used in other cell types and organisms to mediate processes that require cytoplasmic exclusion or might function as a barrier to create cytoplasmic regions with different compositions.
Several aspects of our results seem particularly interesting and perhaps unexpected. First, movement of the cones relies on the bundles, not the meshwork. In other motility systems, such as the leading edge of motile cells and Listeria comet tails, Arp2/3 complex–mediated meshwork formation is thought to be important for motility (Carlier and Pantaloni, 2007). In the case of actin cones, the meshwork forms just as movement initiates. Thus, we expected that the meshwork would be critical for movement. However, cones that lack meshwork (arp3 mutant) can move.
Second, profilin loss of function affects the formation of bundles but not meshwork. Profilin has been implicated in both formin-mediated actin bundle formation and Arp2/3 complex–mediated meshwork formation. Therefore, we might have expected both meshwork and bundle formation to be affected by profilin loss of function. During Arp2/3 complex–dependent Listeria monocytogenes motility in vitro, profilin is not absolutely required, but accelerates the rate of movement (Loisel et al., 1999). Perhaps the slow turnover of filaments in the cones (Noguchi and Miller, 2003) compared with turnover in Listeria comet tails (Theriot et al., 1992) reduces the need for profilin-mediated acceleration of monomer addition. Alternatively, maybe the small amount of activity that remains in this hypomorphic mutant is sufficient to participate in actin cone meshwork formation. Another possible explanation is that a different regulator of assembly, such as capulet (Baum et al., 2000; Baum and Perrimon, 2001), a CAP/SRV2 ortholog, or ciboulot (Boquet et al., 2000), a thymosin β-4 ortholog, has a more important role in regulating monomer availability and addition during actin cone meshwork formation.
Interestingly, bundle formation has a stringent requirement for profilin. Although we have not yet been able to directly implicate a formin in bundle formation (see below), it seems likely that actin cone bundles would be nucleated by a formin, similar to bundles in other cells (Evangelista et al., 2002, 2003; Sagot et al., 2002). The fact that profilin activity is required suggests that the profilin-mediated gating of monomer addition during formin nucleation that has been observed in vitro (Kovar et al., 2003, 2006; Romero et al., 2004) is likely to be critical in vivo. This is similar to the situation in yeast where the formin, Bni1, and profilin are both required for and work together during actin cable assembly (Evangelista et al., 2002; Sagot et al., 2002).
The protein responsible for nucleating bundle formation remains unidentified, although a formin seems the most likely candidate. There are six predicted formin orthologues in Drosophila (Goldstein and Gunawardena, 2000; Higgs and Peterson, 2005). We have examined the diaphanous mutant (Castrillon and Wasserman, 1994), but defects that occur during the spermatocyte divisions make interpretation of phenotypes during individualization difficult (T. Noguchi, unpublished observations). Of the other five predicted formin orthologues in Drosophila, two (CG6807, GC5797) are not yet characterized and mutants are not available at present. Mutations have been identified in the other formins, cappuccino (Manseau and Schupbach, 1989; Emmons et al., 1995), DAAM (Matusek et al., 2006), and formin3 (Tanaka et al., 2004). cappuccino mutations do not appear to affect spermatogenesis, whereas DAAM and formin3 mutations are lethal. Further experiments will be required to determine which nucleator is important here.
In studies of the interplay between actin bundles and meshwork at the leading edge of motile cells, bundled filaments of filopodia were hypothesized to arise via convergence of dendritic arrays of actin in lamellipodial meshwork (Svitkina et al., 2003). However, this idea has been challenged by the demonstration that in both mammalian cells and Dictyostelium, filopodia still form when Arp2/3 complex–mediated meshwork assembly is absent (Steffen et al., 2006). Similarly, in actin cones, bundle formation is not dependant on the meshwork. Bundles form before the meshwork is present and can form even when meshwork never forms, as in the arp3 mutant. In addition, the meshwork does not appear to be a precursor for the bundles during movement, because there is no "flow" of actin from the meshwork into the bundles (Noguchi and Miller, 2003).
An open question is what provides the force for movement. Monomer addition at the barbed ends of the bundles could propel the cone away from the membrane, which would result in movement in the forward (pointed end) direction. Actin turnover (assembly and disassembly) is required for movement (Noguchi and Miller, 2003), but whether this is important for force production remains unclear. If barbed-end elongation of bundles mediates movement, we would expect to see movement of a bleached spot of GFP-actin from the rear to the front of the cone in photobleaching experiments. However, we did not detect any movement of bleached areas, either forward or backward, in FRAP experiments (Noguchi and Miller, 2003). Motors may be involved, although it is unlikely that myosin VI motor activity is required. The cones move at normal speed (at least in the initial stages) when myosin VI function is absent (Noguchi et al., 2006). We cannot eliminate a contribution by another myosin, perhaps binding to the axoneme using its tail and moving in the barbed-end direction, allowing the observed cone movement in the pointed-end direction. The only other myosin that is known to play a role in spermatogenesis is myosin V (Mermall et al., 2005). In myosin V mutant testes, actin cones do not form normally and no normal individualization complexes can be seen moving along the axonemes. These abnormalities preclude drawing conclusions about whether myosin V has a role in cone movement. Alternatively, a microtubule-based motor that uses the axoneme as a track and pulls the cones forward remains a possibility. However, why actin turnover would be important for either microtubule or actin motors to move the cones is unclear. Further study will be required to fully understand all the interesting aspects of this structure's formation, regulation, and motility.
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ACKNOWLEDGMENTS |
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Footnotes |
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These authors contributed equally to this work. 
|| Present address: Consortium for Policy Research in Education, University of Pennsylvania, 3440 Market Street, Suite 560, Philadelphia, PA 19104-3325.
Address correspondence to: Kathryn G. Miller (miller{at}wustl.edu)
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