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Vol. 19, Issue 10, 4287-4297, October 2008
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Faculty of Life Sciences, Wellcome Trust Centre of Cell-Matrix Research, Manchester M13 9PT, United Kingdom
Submitted February 20, 2008;
Revised July 16, 2008;
Accepted July 18, 2008
Monitoring Editor: Josephine C. Adams
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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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), and, accordingly, muscle-attached hypodermal cells in Caenorhabditis elegans are highly enriched in intermediate filaments, as is similarly the case for keratinocytes in the mammalian skin. Failure of such intermediate filaments to form or anchor appropriately, has dire effects on the cells' integrity (Jonkman, 1999; Bosher et al., 2003). In Drosophila, intermediate filaments are virtually absent (Bartnik and Weber, 1989; Adams et al., 2000; Goldstein and Gunawardena, 2000), and alternative cytoskeletal solutions have to be in place. In epidermal muscle attachment cells of Drosophila (hereafter referred to as tendon cells), the solution to this problem is the development of dense arrays of microtubules. The minus ends of these microtubules are anchored via apical cell junctions to the exoskeletal matrix (cuticle), and their plus ends via integrin-dependent basal cell junctions to tendon matrix. This tendon matrix is also associated with the tips of muscles, thus constituting the myo-epidermal link (Reedy and Beall, 1993; Prokop et al., 1998a,b; Tucker et al., 2004; Bökel et al., 2005).
During the last few decades, Drosophila tendon cells have become exciting model systems to be used for various venues of research. For example, in the context of tendon cells, complex signaling events have been unraveled (Volk, 1999, 2006). Tendon cells provide spatial cues attracting motile tips of developing muscles. Conversely, these muscles reinforce differentiation of the targeted tendon cells. Furthermore, many cellular assembly processes have been addressed, including the differentiation of cell junctions on basal and apical cell surfaces, as well as the establishment of the cytoskeletal architecture and its molecular links to the cell surface (e.g., Prokop et al., 1998a,b; Tucker et al., 2004; Bökel et al., 2005). Apart from integrins, several components have been identified, all of them known to be important in mammalian cell biology. Among these are integrin-linked kinase (Ilk; Zervas et al., 2001), focal adhesion kinase (FAK; Palmer et al., 1999), Paxillin (Yagi et al., 2001), PINCH (Clark et al., 2003), Talin (Brown et al., 2002), Tensin (Blistery) (Torgler et al., 2004), and the BPAG1/ACF7 orthologue Short stop (Röper et al., 2002).
Apart from providing an excellent model system for the study of cell signaling and assembly processes, tendon cells may also provide a representative model for certain cell types in vertebrates. An example of this is the similarity between Drosophila tendon cells and the Deiter's and pillar cells in the organ of Corti of the inner ear. These cells provide rigidity and support for sensory hair cells so that they are able to pivot when deflected by basilar membrane movement relative to the tectorial membrane (Forge and Wright, 2002). Hence, these cells also function in an environment of considerable mechanical challenge. Most interestingly, they share some of the properties of Drosophila tendon cells: 1) they contain apico-basal arrays of microtubules (Forge and Wright, 2002), 2) these microtubules display the unusual diameter of 15 rather than 13 protofilaments (Saito and Hama, 1982; Mogensen and Tucker, 1990), and 3) both of them are rich in Spectraplakin proteins (mammalian BPAG1 vs. its close Drosophila orthologue Short stop, Shot) (Leonova and Lomax, 2002; Röper et al., 2002). This suggests that tendon cells may not be an insect-specific cell type, but they have structurally related counterparts in vertebrates. However, in contrast to current descriptions of tendon epithelial cells, Deiter's and pillar cells have been reported to contain arrays of anti-parallel actin filaments lying interspersed with the microtubule arrays, both of which are heavily cross-linked (Slepecky and Chamberlain, 1983; Arima et al., 1986). These actin fibers have been suggested to convey stability rather than dynamic morphogenetic properties to the support cells (Slepecky and Chamberlain, 1987).
Here, we show that prominent F-actin arrays also exist in Drosophila tendon cells. Based on refined imaging approaches and a number of molecular markers we show that these fibers project apico-basally, fully overlap with the microtubule arrays, and are associated with type II myosin. Our data suggest that the F-actin arrays of tendon cells are additional architectural elements of tendon cells, but they display properties very distinct from stress fibers.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), anti-mCD8 (rat, 1:10; Invitrogen, Carlsbad, CA), anti-Disk large (Dlg) (rabbit, 1:1000; courtesy of U. Thomas, Institute for Neurobiology, Magdeburg, Germany; Woods and Bryant, 1991), anti-βPS-integrin (CF6G11 ascites; mouse, 1:300; courtesy of E. Martin-Blanco, Instituto de Biologia Molecular de Barcelona, Barcelona, Spain; Brower et al., 1984); FAK (rabbit, 1:1000; courtesy of R. H. Palmer, Umea University, Umea, Sweden; Grabbe et al., 2004); anti-c-Paxillin (catalog no. 610051; mouse, 1:100; BD Biosciences Transduction Laboratories, Lexington, KY); anti-
-actinin (MAC276 ascites; rat, 1:10; courtesy of B. Bullard; Lakey et al., 1990); anti-Enabled (5G2; mouse, 1:20; Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA), anti-tubulin (mouse, 1:1000; T-9026; Sigma Chemical. Poole, Dorset, United Kingdom), anti-tyrosinated tubulin (MAB1864; rat, 1:500; Millipore Bioscience Research Reagents, Temecula, CA), anti-glu tubulin (rabbit, 1:200; Millipore Bioscience Research Reagents), anti-Short stop (guinea pig; 1:100; courtesy of T. Volk; Strumpf and Volk, 1998), and preabsorbed secondary antibodies (donkey; 1:200; Jackson ImmunoResearch, West Grove, PA). Cyanine (Cy)3- or Cy5-coupled phalloidin was diluted 1:500 or 1:100, respectively, and applied for 1 h.
Dissection of late stage 17 embryos (stages according to Campos-Ortega and Hartenstein, 1997) or of late L3 larvae was carried out in phosphate-buffered saline (PBS) followed by 1-h fixation in 4% paraformaldehyde and 1 h wash in PBS plus 0.3% Triton X-100, with the following two exceptions: anti--actinin and anti-c-paxillin stainings were carried out on larvae fixed for 1 min in 60°C PBS, and EB1::green fluorescent protein (GFP)-expressing animals were treated for 4 min in –20°C methanol. For live imaging (Figure 7), embryos were manually devitellinized and mounted in PBS under a coverslip. Imaging was carried out with an AxioCam (Carl Zeiss, Jena, Germany) on a BX50WI fluorescent microscope (Olympus, Tokyo, Japan) or with an SP5 confocal microscope (Leica, Wetzlar, Germany).
Fly Stocks
The following fly stocks were used: UAS-GFP-a-tub84B (courtesy of C. Boekel, Dresden Technical University, Dresden, Germany; Grieder et al., 2000), UAS-ena::GFP (courtesy of T. Millard; Gates et al., 2007), UAS-Shot-PA::GFP (courtesy of P. Kolodziej, Vanderbilt University Medical Center, Nashville, TN, deceased; Lee and Kolodziej, 2002), UAS-EB1::GFP (courtesy of P. Kolodziej; unpublished), UAS-actin::GFP (UAS-Act5C.T:GFP127.37.2 and UAS-Act5C.T:GFP127.18.4; Bloomington Stock Center, Indiana University, Bloomington, IN; Kelso et al., 2002), UAS-mCD8-GFP (courtesy of L. Luo, Stanford University, Stanford, CA; Lee and Luo, 1999), sqhAX3; P[w+ sqh-gfp]42 in which the transgene is the only source of regulatory light chain (courtesy of E. Martin-Blanco; Royou et al., 2002), UAS-Rho1-dsRNA8.2, UAS-Rho1-dsRNA8.1/TM3,Sb1, UAS-Rho1.N192.1 (Bloomington Stock Center), shotsf20 (Prokop et al., 1998b); shot3 (courtesy of P. Kolodziej; Lee et al., 2000), stripe-Gal4 (courtesy of T. Volk; Subramanian et al., 2003), Neurexin IV::GFP (line CA06597 obtained from FlyTrap), myospheroid (mysXG43; courtesy of N. Brown, University of Cambridge, Cambridge, United Kingdom; Bunch et al., 1992), UAS-shot-dsRNA34 (courtesy of T. Volk, Weizmann Institute, Rehovot, Israel; Subramanian et al., 2003), integrin-linked kinase::GFP (IlkZCL3111; FlyTrap line, Bloomington Stock Center), collagen IV::GFP (vkgG454; FlyTrap line G00454; courtesy of E. Martin-Blanco; Morin et al., 2001), and enaGC1 and ena23 (Bloomington Stock Center; Wills et al., 1999).
Application of Toxins
Toxins were applied in PBS to cultured fillet preparations of late stage 17 or late L3 larvae in PBS or commercial Schneider's medium (Invitrogen) by using the following concentrations and incubation times: 1–2.5 h in 1–100 µM nocodazole (made up from a 5 mM stock in dimethyl sulfoxide [DMSO]); 1 mg/ml collagenase IV (Sigma Chemical) for 5–8 min (muscle detachment monitored under dissection microscope); 30 min in commercial trypsin-EDTA 1x solution for cell culture (Lonza 17-161, Lonza Verviers SPRL, Verviers, Belgium; 500 mg/l trypsin); 1–2.5 h in 4 µM cytochalasin D (made up from a frozen 2 mM stock solution in DMSO) and/or 6 µM latrunculin A; and 1 h in 50 µM or 1 mmol of Y-27632 (Sigma Chemical; made up from a 29 mM stock in water; NIH3T3 fibroblast cultured were treated for 1 h in 30 µM Y-27632 in DMEM medium). Collagenase IV or trypsin treatments were followed by at least 15 min in PBS or Schneider's medium.
Electron Microscopy
Electron microscopy was performed as described previously (Budnik and Ruiz-Cañada, 2006). For the ultrastructural analyses of collagenase IV-treated animals, St.17/L1 animals were filleted on Sylgard as described previously (Budnik and Ruiz-Cañada, 2006), treated for 5–6 min in 1 mg/ml collagenase IV (Sigma Chemical) in saline, briefly washed in pure saline, and then fixed for 1 h in 2.5% GDA in 0.05 M phosphate buffer, followed by further standard procedures (Budnik and Ruiz-Cañada, 2006).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Because microtubules are the most prominent structural feature of tendon cells (see Introduction), we started our light microscopic analyses with targeted expression of UAS-tubulin::GFP by using the tendon cell-specific driver line stripe-Gal4 (Gal4/UAS-system described in Duffy, 2002). At indirect muscle attachments, fluorescence is arranged into speckled or dashed fields of 30–40 µm in diameter, which we interpret as primarily vertically oriented arrays of microtubules (Supplemental Figure 1F). At direct muscle attachments, similar speckled or dashed fields also exist (Figure 1, B and C4). However, if the muscles are sufficiently stretched, part of these fibers is arranged into dense fluorescent bands measuring 2.5 to 5 µm in width, depending on the level of stretch. We interpret these bands (hereafter referred to as "cytoskeletal belts") as oblique microtubules arranged in parallel to the axis of the pulling muscles (between arrow heads in Figures 1 and 2). This interpretation of the cytoskeletal arrays is in full agreement with all further observations made during this project.
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). All of these markers show enrichment at the cytoskeletal belt (Figure 2, E–H) and, among them, Shot displays a more elaborate pattern within the cytoskeletal belt: two prominent bands of protein accumulation run in parallel to the muscle tip on the proximal and distal side of the cytoskeletal belt. In between these bands, Shot expression is lower and often occurs in fiber-like arrangements (Figure 2, F and G).
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), it could be acting here to stabilize the microtubule arrays in tendon cells. Second, the cytoskeletal belt can be spatially subdivided, suggesting that the two bands of prominent Shot::GFP localization correspond to the apical and basal ends of the microtubule arrays. This would mean that apical and basal surfaces of tendon cells can be distinguished at the light microscopic level, with obvious implications for future studies. The following experiments were performed to test these two hypotheses.
Tubulin Arrays Are Highly Stable and Can Be Resolved in the Apicobasal Axis
To test for a potential stability of microtubules in tendon cells, we cultured fillet preparations of larvae for 2–2.5 h in medium containing high concentrations (10–100 µM) of nocodazole. Nonarray microtubules in these tendon cells are vastly eliminated, demonstrating the strength of the treatment (Figure 2B). In contrast, the microtubule arrays are fully protected (Figure 2B'). Thus, the arrays contain stable microtubules, and Shot is a potential factor mediating this property.
To explore the apicobasal resolution of the cytoskeletal belts, we studied the localization of known marker molecules by using either antibodies or fly lines carrying GFP-tagged versions of endogenous proteins. Neurexin IV localizes to septate junctions along the lateral side of epidermal cells (Baumgärtner et al., 1996; Knust, 2000). Accordingly, Neurexin IV::GFP delineates all epidermal cells, including the tendon cells. Analyses of such specimens reveal that, in contrast to the planar arrangement of most other epidermal cells, tendon cells take on a step-like shape in sagittal view (Figure 1, B and D2), most likely imposed by the attached muscle. When viewed as a frontal section through a confocal image stack (Figure 1D1), the cytoskeletal belt (visualized with phalloidin, as explained in the next chapter) is framed on each side by lines of Neurexin IV::GFP expression (Figure 1D1). Similar observations were made with anti-Dlg staining, a further marker for septate junctions (Knust, 2000; Figure 1A). These observations clearly demonstrate that the cytoskeletal belts span tendon cells in an apico-basal direction. To confirm our finding further, we probed marker molecules known to be associated with myotendinous junctions, i.e., expected to localize to the basal end of the cytoskeletal arrays. Indeed, stainings for βPS-integrin, Ilk::GFP, FAK, or Paxillin all localized to the basal end of the cytoskeletal arrays (shown for Ilk; Figure 3C1).
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). We tested this aspect at the confocal level using a GFP-tagged version of the integral basement membrane component collagen IV (Viking::GFP). The Viking::GFP localization pattern clearly confirmed the exclusion of basement membrane from myotendinous junctions (arrows in Figure 1E), further demonstrating the spatial resolution of the confocal analysis and its precise correlation with ultrastructural findings. In summary, we have refined strategies which enable us to carry out more detailed microscopic analyses of tendon cells, resolving cytoskeletal bands in the apico-basal axis and resolving the tendon cell versus muscle side at muscle attachments. Applying these strategies, we investigated further properties of tendon cells.
Tendon Cells Contain Prominent Arrays of Stable F-Actin
Support cells in the Organ of Corti show arrays of highly cross-linked actin filaments, intermingled with the prominent arrays of microtubules (Slepecky and Chamberlain, 1983; Arima et al., 1986). This tempted us to investigate the presence of similar structures in tendon cells. To this end, we targeted expression of actin::GFP to these cells. To our surprise, actin::GFP is strongly expressed in all areas of the cytoskeletal arrays and forms a fibrous cytoskeletal belt (Figure 5A3), very similar to that seen for tubulin::GFP (compare Figures 5A and 4E vs. 3A and 2A2). The only difference is that actin, but not tubulin, has a tendency to enrich apically and basally, most likely due to its incorporation into submembranous densities of hemi-adherens junctions at the apical and basal tendon cell surfaces (Figure 4, D and E). The current assumption, that microtubules are the sole stabilizing elements of tendon cells, needs to be reconsidered.
The phalloidin staining pattern in tendon cells is congruent to the localization pattern of actin::GFP (Figures 4E and 5A). This supports our findings and provides a simple labeling strategy for further experimentation. In double labeling experiments, phalloidin marks the same area as tubulin::GFP (Figure 1C3), and this becomes especially obvious upon muscle detachment (Figures 3B3 and 5F). Therefore, we propose that F-actin and microtubule arrays are intermingled. Such a scenario resembles the parallel arrangement of actin filaments and microtubules demonstrated previously for epidermal cells of developing Drosophila wings (Mogensen and Tucker, 1988). These wing epithelial cells share a variety of structural features with tendon cells and likewise have to withstand enormous mechanical challenges, especially when the wing blades unfold in the freshly hatched fly (Bökel et al., 2005).
Ultrastructural studies using S1 decoration in support cells showed actin fibers to be anti-parallel (Slepecky and Chamberlain, 1983; Arima et al., 1986). To address this property, we monitored tendon cells for localization of Enabled, a molecule of the actin assembly machinery binding to barbed ends of F-actin (Krause et al., 2003). Although the staining for anti-Enabled was enriched in late larval tendon cells, compared with the surrounding epidermal cells, there was no obvious enrichment at the cytoskeletal arrays (Figure 6C). This was likewise the case when targeting Ena::GFP to late larval tendon cells (Figure 6D). In late embryos, Enabled staining was often enriched in dots along the lateral border of tendon cells, but it was never associated with the cytoskeletal belts (Figure 6E). Thus, these experiments do not provide any insights into the orientation of F-actin in tendon cells. However, they suggest that these fibers display low dynamism and might therefore not depend on an efficient F-actin polymerization machinery, as seen in other F-actin–rich structures such as filopodia of growth cones (Dwivedy et al., 2007; our unpublished results for Drosophila). In agreement with this notion, enaGC1/ena23 loss-of-function mutant embryos, which lack prominent Enabled expression in tendon cells (Figure 6F) and display severe CNS phenotypes (data not shown), fail to display obvious phenotypes of the F-actin arrays (Figure 6H). Low dynamism of F-actin arrays may imply that they are stable. We tested this possibility by treating tendon cells with F-actin destabilizing drugs. Such treatment causes actin to aggregate into dots dispersed throughout tendon cells, demonstrating the effectiveness of the treatment, whereas the F-actin arrays were fairly unaffected (Figure 5B). Even 2.5-h incubation with a cocktail of 4 µM cytochalasin D together with 6 µM latrunculin A, failed to abolish the F-actin arrays (data not shown). This resistance to destabilizing drugs is in agreement with the observation that the apicobasal actin fiber arrays in support cells of the Organ of Corti are stable (Slepecky and Chamberlain, 1987; Oshima et al., 1992).
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). Stress fibers contain -Actinin and nonmuscle Myosin type II (Lo et al., 2004; Lele and Kumar, 2007). Support cells in the Organ of Corti also seem to contain these proteins (Drenckhahn et al., 1982; Slepecky and Chamberlain, 1985; Donaudy et al., 2004). Therefore, we investigated the presence of these factors in Drosophila tendon cells.
Antisera against Drosophila -actinin reveal strong staining in Z-discs of the muscle and at the myotendinous junction (data not shown), but no -actinin was detectable on the cytoskeletal arrays of tendon cells or on their basal surfaces upon muscle detachment (data not shown). However, on the basis of these results, the presence of -actinins in tendon cells cannot be fully excluded because a paralogue, -actinin3, is reported in FlyBase (Drysdale et al., 2005), for which no antibody is available.
To address the presence of myosin II, we used a previously published fly line expressing Sqh::GFP (a GFP-tagged version of type II myosin regulatory light chain, Spaghetti squash) under the control of the sqh promoter in a sqh mutant background (Royou et al., 2002). In these larvae, only tendon cells, but not any other epidermal cells, showed cytoplasmic enrichment of Sqh::GFP to various degrees (data not shown). Most importantly, most tendon cells show prominent localization of Sqh::GFP to the cytoskeletal arrays (Figure 6, A1 and A2). We confirmed this finding by staining against endogenous Zipper protein, the nonmuscle Myosin II heavy chain (Young et al., 1993). Anti-Zipper staining was enriched particularly in tendon cells, in which it colocalizes with Sqh::GFP at the cytoskeletal belts (Figure 6B).
Arrays of F-Actin in Tendon Cells Form in the Absence of Rho1, Rock, or Integrin Activity
Encouraged by our finding with myosin II, we tested whether essential regulators of stress fibers, such as Rho1, its effector Rock, and integrins (Schoenwaelder and Burridge, 1999), are involved in the regulation of F-actin arrays in tendon cells. First, we down-regulated Rho1 activity via tendon cell-specific expression of Rho1-specific antisense RNA or of Rho1N19, a dominant-negative version of Rho1. Both treatments caused lethality at the late L2/early L3 larval stage. However, in contrast to ordinary stress fibers in migrating cells (Schoenwaelder and Burridge, 1999), the cytoskeletal belts in such tendon cells clearly failed to vanish upon inhibition of Rho1 activity (Figure 7, B and C). We complemented these studies with experiments using Y-27632, a selective inhibitor for Rock. Rho1 is generally believed to activate myosin light chain through Rock (Govek et al., 2005), and this holds true in Drosophila (Verdier et al., 2006). Previously, Y-27632 has been successfully used in Drosophila and was shown to have a direct effect on Sqh::GFP localization (Royou et al., 2002). However, in our experiments, one hour incubations in Y-27632 at conventional or elevated doses (50 µM or 1 mmol) did not affect F-actin arrays in tendon cells or the localization of Sqh::GFP (data not shown), whereas parallel treatment of mouse NIH3T3 fibroblasts for the same period completely disassembled their stress fibers (data not shown).
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). In mysXG43 mutant embryos expressing actin::GFP in tendon cells, potential F-actin arrays can be seen in only a subset of tendon cells where they occur in unusual ring structures (Figure 7D'). We observed the same phenomenon when using tubulin::GFP expression (Figure 7E'). At first sight, this seemed to contradict previous ultrastructural analyses showing that microtubules are present in mys mutant tendon cells. However, those electron microscopy studies also showed that microtubule arrays are dramatically shortened in mysXG43 mutant tendon cells (Prokop et al., 1998a). We therefore reasoned that subdetectable levels of actin::GFP or tubulin::GFP in many mysXG43 mutant tendon cells might be due to severe shortening, rather than the absence of the cytoskeletal arrays. In contrast to this finding, we predicted that molecular components of the myotendinous junction, such as Ilk::GFP, should be abundantly present in mys mutant embryos, based on the former report that hemiadherens junctions in mys mutant tendon cells are unaffected at the ultrastructural level (Prokop et al., 1998a). Indeed, Ilk::GFP was found to be reliably expressed in mysXG43 mutant tendon cells, where it forms the same ring structures as actin::GFP and tubulin::GFP (Figure 7F'). Hence, our light microscopic analyses are in accordance with previous electron microscopic studies, and we propose that only the length and shape, but not the principal formation, of F-actin arrays is dependent on integrin function.
In fibroblasts, stress fibers are positively regulated through mechanical force, partly mediated through type II myosin function (Bershadsky et al., 2006; Lele and Kumar, 2007). Such mechanical forces are abolished in mysXG43 mutant embryos where muscles detach from tendon cells (Prokop et al., 1998a). Therefore, the F-actin arrays we found to persist in mysXG43 mutant embryos suggest that mechanical force is not crucial for their formation. To test this hypothesis more directly, we detached muscles from tendon cells at the late larval stage via collagenase IV- or trypsin treatment and cultured these specimens for 2–3 h after detachment. However, under these conditions, neither intensity nor distribution of actin::GFP, tubulin::GFP, or Sqh::GFP seemed gravely affected (actin::GFP shown in Figure 5D). Application of 4 µM cytochalasin D during the culturing period did not suggest any noticeable increase in the vulnerability of the actin fibers (Figure 5E). These findings are consistent with our studies in integrin-deficient embryos.
We conclude from these studies that the F-actin fibers in tendon cells fail to display regulatory properties described for stress fibers, and the role of type II myosin in these fibers remains elusive.
F-Actin Arrays Maintain a Physical Apico-Basal Link When Microtubule Arrays Are Dysfunctional
We next tested the properties of F-actin arrays under conditions when the microtubule arrays are affected. For example, the actin–microtubule linker Shot of the Spectraplakin family is required for the formation, and/or maintenance, of the microtubule arrays in tendon cells (Gregory and Brown, 1998; Prokop et al., 1998b; Strumpf and Volk, 1998; Subramanian et al., 2003). We therefore analyzed the presence and stability of F-actin arrays upon loss of Shot function in tendon cells. When knocking down shot with a previously published interference RNA (iRNA) construct, we found significant elongation of tendon cells and their cytoskeletal belts (Figure 8C), as reported previously (Subramanian et al., 2003). F-actin arrays of such embryos treated for 1 h with 4 µM cytochalasin D, or with PBS as a control, were very similar in appearance (Figure 8D). Because knockdown in these animals might not be complete, we combined iRNA expression with one copy of the shot3 null allele or analyzed shotsf20/shot3 mutant embryos (the strongest known alleles). In either case, muscle tips are maximally retracted from the cuticle. However, in spite of this dramatic elongation of tendon cells, long thick F-actin cables maintain a link between cuticle and the severely retracted muscle tips (Figure 8, E–J). Expression of tubulin is very low in these structures (data not shown), in agreement with previous reports that tubulin levels are reduced in shot mutant embryos (Strumpf and Volk, 1998). When challenging the F-actin cables with cytochalasin D, latrunculin A, or a combination of both agents, treated and sham-treated embryos did not reveal any obvious differences (Figure 8, E–J).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, 2006). Here, we have delivered refined light and laser microscopic approaches providing enhanced opportunities for future work in this system. Applying these strategies, we have refined insight into the organization of tendon cells, and we have uncovered new insight into their molecular composition.
The essential advance in microscopic analyses is the clear demonstration that tendon cells can be resolved in the apicobasal axis at the position of the cytoskeletal belt and that subcellular markers, such as Paxillin, Ilk, integrin, or Neurexin IV, can be mapped along this axis. The proteinase-based detachment assay provides a quick means to confirm potential localization of proteins to basal surfaces of tendon cells. By showing that apical and basal ends of cytoskeletal belts can be distinguished, these prominent structures have become helpful landmarks in their own right, which can be easily visualized via phalloidin staining. From the studied markers in conjunction with new ultrastructural data, we are confident that our model of the subcellular organization of tendon cells at the light microscopic level (Figure 1B) provides a reliable readout with good spatial resolution. Via these strategies, it is now possible to assign proteins rapidly to subcellular compartments, making us less dependent on complex immunoelectron microscopic localization studies. For example, by using a GFP-tagged version of endogenous collagen IV (Vkg), we could establish effortlessly that basement membrane components are absent from the myotendinous junction (Figure 1E), whereas previously we were unable to answer this question unequivocally using detailed ultrastructural studies (Prokop et al., 1998a; arrows in Figure 4).
Shared Features between Tendon Cells and Support Cells in the Inner Ear of Vertebrates
In addition to providing refined descriptions of the organization of tendon cells, we have reported new molecular features for this cell type. Their microtubular arrays are resistant to nocodazole, and they display prominent arrays of stable F-actin that seem to be associated with type II myosin (Sqh and Zipper). These observations extend the catalogue of properties shared between Drosophila tendon cells and support cells in the inner ear of vertebrates. Both display apico-basal arrays of microtubules, which are stable and contain the unusual number of 15 protofilaments (Slepecky et al., 1995), both display apicobasal arrays of stable actin filaments, and both seem to express type II myosin and Spectraplakins. This leads us to speculate that the organization of both cell types could be analogous or even homologous, and work in tendon cells might become a source of inspiration with potential translational value, e.g., in the context of age-related deafness caused by decay of support cells (Saha and Slepecky, 2000; Forge and Wright, 2002). However, it needs to be pointed out that -actinin, which is present in support cells of the inner ear, might be absent in Drosophila tendon cells.
Shared and Distinct Features of Hemi-adherens Junctions and Focal Adhesions
The existence of F-actin arrays in tendon cells sheds new light into focal adhesion components which are prominently expressed at the basal surface of tendon cells (see Introduction and Figure 3). Such components are best understood in migrating cells in culture, in which they assemble into integrin-dependent membrane-associated complexes called focal adhesions (Zaidel-Bar et al., 2004). Focal adhesions anchor and regulate actin stress fibers, contractile cell-spanning structures associated with factors such as type II myosin and -actinin (Lo et al., 2004; Lele and Kumar, 2007). In addition, focal adhesions are also targeted by microtubules (Bershadsky et al., 2006), the second prominent cytoskeletal component attached to myotendinous junctions in tendon cells. However, there are clear differences between both adhesion complexes. Loss of integrin function correlates with loss of focal adhesions and stress fibers (Schoenwaelder and Burridge, 1999). In contrast, the hemi-adherens junctional complex of tendon cells and cytoskeletal arrays are, in principle, formed and maintained in the absence of integrins and muscle adhesion, as demonstrated ultrastructurally elsewhere (Prokop et al., 1998a) and here by the maintained localization of various markers, such as actin, tubulin, and the focal adhesion component Ilk (Figure 7, D–F'). Down-regulation of Rho or Rock is known to reduce focal adhesions and stress fibers (Schoenwaelder and Burridge, 1999), which does not hold true for F-actin arrays in tendon cells (Figure 7, A–C). Similarly, microtubules negatively influence focal adhesions (Bershadsky et al., 2006), whereas they are stably anchored at the myotendinous junctional complex of tendon cells. Thus, although both adhesion complexes contain similar molecular players, their systemic output is clearly distinct. It will be very interesting to carry out comparative studies of these proteins to appreciate the breadth of their functions in the context of cell migration versus integrity. For example, it might not be unrealistic to speculate that a simple molecular switch, such as Vinculin (Humphries et al., 2007), could turn a Rho-dependent into a Rho-independent constellation. The strategies and F-actin arrays reported here provide an improved experimental platform for such analyses in tendon cells.
The Potential Role of F-Actin Arrays
A very likely role for F-actin arrays is the provision of additional stability, enabling tendon cells to withstand forces built up between contracting muscles and the rigid cuticle. In general, microtubules are ideal architectural elements to provide stability against shear forces (Howard, 2001), an obvious challenge tendon cells are exposed to. Microtubules are strong in resisting tensile force. However, they are likely to be intolerant to overstretching, and they would be expected to break when elongated >1% of their length (Howard, personal communication). The F-actin arrays may be an ideal complementary system in this situation. Their architectural importance is best demonstrated in situations where microtubule arrays are significantly impaired, as is the case in shot mutant embryos (Figure 8). Initially, the enormous lengthening of F-actin observed under these conditions seems surprising given that actin filaments tend to break when overstretched, even more when under torsion (Tsuda et al., 1996). However, F-actin in cellular networks displays vastly enhanced elastic properties through cross-linking factors and myosin association (Gardel et al., 2006). Furthermore, length adaptation through active polymerization might prevent a force overload of the array system.
Actin fibers might also play a role during development. Thus, work in cultured tendon cells has suggested that microtubules in developing tendon cells nucleate at the apical surface and extend toward the basal side where their plus ends are precisely targeted to pre-existing cortical capturing sites provided by the hemi-adherens junction (Tucker et al., 2004). Potentially pre-existing actin fibers could guide these microtubule targeting events by serving as tracks for their plus ends. A similar scenario has been proposed for the targeting of microtubules to focal adhesions (Kodama et al., 2004). However, this possibility can only be tested once we understand the mechanisms underlying F-actin array formation.
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ACKNOWLEDGMENTS |
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Footnotes |
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* These authors contributed equally to this work. 
Present addresses: 
Program Unit Development and Genetics, Laboratory for Molecular Developmental Biology, LIMES-Institute, University of Bonn, D-53115 Bonn, Germany;
Department of Craniofacial Development, King's College, Guy's Hospital, London SE1 9RT, United Kingdom.
Address correspondence to: Andreas Prokop (andreas.prokop{at}manchester.ac.uk).
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