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Keywords:

  • Drosophila;
  • dorsal closure;
  • JNK;
  • integrin;
  • myospheroid;
  • scab

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

Epithelial movements are key morphogenetic events in animal development. They are driven by multiple mechanisms, including signal-dependent changes in cytoskeletal organization and in cell adhesion. Such processes must be controlled precisely and coordinated to accurately sculpt the three-dimensional form of the developing organism. By observing the Drosophila epidermis during embryonic development using confocal time-lapse microscopy, we have investigated how signaling through the Jun-N-terminal kinase (JNK) pathway governs the tissue sheet movements that result in dorsal closure (DC). We find that JNK controls the polymerization of actin into a cable at the epidermal leading edge as previously suggested, as well as the joining (zipping) of the contralateral epithelial cell sheets. Here, we show that zipping is mediated by regulation of the integrins myospheroid and scab. Our data demonstrate that JNK signaling regulates a set of target genes that cooperate to facilitate epithelial movement and closure. Developmental Dynamics 235:427–434, 2006. © 2005 Wiley-Liss, Inc.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

Cell migration, either by individual cells or by cell groups, is observed as part of a multitude of normal and pathologic processes, including development, wound healing, immune response and cancer (reviewed in Ridley et al., 2003). Although migration is seen in many different cell types, a basic set of features is conserved. Cells are typically polarized toward the direction of movement and exhibit a contractile actin-based apparatus localized at the leading edge, which produces the force necessary for movement. Integrins mediate controlled adhesion to the extracellular matrix (ECM) and, in this manner, contribute to the regulation of cellular movements. The coordination of contraction/relaxation coupled with attachment/detachment to ECM is regulated by a complex network of signaling interactions. A complete understanding of the mechanisms that control and execute cell migration would be valuable for combating pathologies such as the metastatic spreading of tumors and inflammatory diseases, where regulation of movement appears to break down.

The JNK pathway is increasingly recognized as a regulator of cell morphogenesis and mobility (Xia and Karin, 2004). In cultured mammalian cells, loss of MEKK1, an upstream activating kinase, inhibits JNK activation and cell migration in response to serum stimulation (Xia et al., 2000). Furthermore, cells transfected with dominant-negative forms of JNK lose their ability to migrate (Huang et al., 2003). It is clear that transcriptional responses to JNK are critically involved in the morphogenetic functions of JNK. Mutants lacking Jun, a transcription factor and downstream effector of JNK signaling, fail to complete cell movement processes that normally occur during development, such as eyelid closure in mice and embryonic dorsal closure (DC) in Drosophila (Hou et al., 1997; Kockel et al., 1997; Riesgo-Escovar and Hafen, 1997; Li et al., 2003; Zenz et al., 2003).

DC is a genetically accessible model to investigate how JNK and its target genes orchestrate the different cell biological aspects of epithelial morphogenetic movements (reviewed in Kockel et al., 2001; Harden, 2002; Jacinto et al., 2002b). DC entails the replacement of an extraembryonic dorsal cell layer, the amnioserosa, by an embryonic tissue, the epithelium, during late embryogenesis. The leading edges of the epithelium on both sides of the embryo, which initially surround the amnioserosa, sweep dorsally over the amnioserosa, and fuse at the dorsal midline. The cells at the leading edge (LE) of the epithelial cell sheet share features with other migrating cells: integrin-based adhesions, a contractile band of actin and myosin (known as “supracellular purse string” or “actin cable”) focused at the apical end of the polarized cells, filopodial projections, and extensive cell shape rearrangements during closure (Jacinto et al., 2000; Kiehart et al., 2000; Brown et al., 2002; Kaltschmidt et al., 2002; Hutson et al., 2003).

JNK/Jun signaling is required for DC and loss-of-function mutations of each pathway component tested (e.g., slipper, hemipterous, basket, jun, and kay, encoding the JNKKK, JNKK, JNK, Jun, and Fos, respectively) fail in DC (reviewed in Kockel et al., 2001; Harden, 2002; Jacinto et al., 2002b). JNK signaling regulates changes in the cortical actin cytoskeleton in leading edge cells of moving epithelial sheets (Ricos et al., 1999; Jacinto et al., 2000; Jasper et al., 2001; Kaltschmidt et al., 2002). We have reported previously that chickadee, a gene encoding the Drosophila profilin homologue, is regulated by JNK and is critical for actin polymerization during dorsal closure (Jasper et al., 2001). Several other JNK-responsive genes with potential functions in tissue organization were identified but not functionally characterized.

In addition to JNK-dependent processes that have been described previously (such as actin polymerization, Ricos et al., 1999; Jasper et al., 2001; Kaltschmidt et al., 2002), we find that JNK signaling is essential for the propagation of epithelial cell–cell adhesion as the two leading edges fuse together (known as “zipping”). Zipping is dependent on integrin function, as mutants lacking expression of the βPS integrin myospheroid (mys) fail in this process (Hutson et al., 2003). We show here that the αPS3 integrin scab (scb) is also a mediator of zipping and that JNK signaling controls zipping by regulating the expression of scb and mys. It emerges that multiple genes cooperate to drive DC and that JNK signaling serves to coordinate their specific morphogenetic functions.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

JNK Signaling in DC

DC is driven by dynamic signal-dependent changes in cell shape, mobility, and adhesion. To investigate whether some or all of these functions might be directed by JNK signaling, we used time-lapse confocal microscopy to observe the DC process in embryos that carry loss-of-function mutations in different components of the JNK signaling pathway, including null alleles or hypomorphic mutations in genes encoding Jun, JNK, and JNKKK (Fig. 1, and Supplementary Videos 1–5, which can be viewed at http://www.interscience.wiley.com/jpages/1058-8388/suppmat). Changes in cellular morphology during dorsal closure were visualized in embryos expressing a moesin–green fluorescent protein (GFP) fusion protein that labels the cortical F-actin cytoskeleton (sGMCA transgene, Kiehart et al., 2000).

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Figure 1. A–E: A sampling of frames from videos depicting embryogenesis at the dorsal closure (DC) stage of wild-type (genotypically a paternal rescue of a jun germline clone; A), jun (germline clone, glc; B), slpr (C), bsk (D), and scb (E) embryos. Each sampling represents roughly 3 hr from each video except scb, which shows 5 hr. The asterisks in A denote each canthus progressing in DC. The white arrows show the lack of zipping in C,D,E. Each frame is artificially color coded as follows: green, epithelium; blue, amnioserosa; red, internal structures. The uncolored raw data are available online in video format (see Supplementary Videos 1–5). For comparing signal intensities between structures and for identifying structures with cellular resolution, refer to the online videos. Note that the slpr video has a high green fluorescent protein (GFP) background fluorescence due to maternal contribution of the GFP balancer used in the parent stock and because the embryo contains only one copy of the sGMCA transgene. Scale bar = 10 μm.

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In wild-type embryos, the actin cable is assembled apically in LE cells before the onset of epithelial stretching. As the cable matures, it begins to contract, thereby transforming the LE front from a scalloped to a straightened appearance (Fig. 1A, Supplementary Video 1, and Kiehart et al., 2000). Once cell stretching initiates (due to contributions of both actin cable and amnioserosal cell contraction; Kiehart et al., 2000; Hutson et al., 2003), the contralateral cell fronts are gradually “zipped” together at the two “canthi” located at the anterior and posterior ends of the dorsal hole (Fig. 1A, asterisks). Zipping results in fusion of the two epithelial cell sheets and exerts a force that contributes to hole closure (Hutson et al., 2003).

Videos of embryos that are homozygous for null alleles of jun or slpr and, thus, deficient for the Jun transcription factor or the relevant upstream JNKK kinase, respectively, show severe defects in DC. The different mutant embryos, as well as embryos lacking both the zygotic and the maternal Jun component (jun germ line clones, GLC, Fig. 1B), display common phenotypic defects (Fig. 1B,C, and Supplementary Videos 2, 3; note that the background fluorescence is higher in the slpr movie than in the others, see legend for Fig. 1 for details). Consistent with previous reports describing JNK signaling-deficient embryos (Ricos et al., 1999; Jasper et al., 2001; Kaltschmidt et al., 2002), the actin cable fails to mature into a pronounced structure in jun mutants (Supplementary Fig. S1). The LE front remains scalloped at a stage of DC when it has become prominent and taut in the wild-type embryos (compare Supplementary Video 1 with Videos 2, 3, and see Fig. S1). Thus, based on previous reports and our data, it is reasonable to hypothesize that JNK signaling is required for efficient formation of the leading edge actin cable. To lend further support to this hypothesis, we asked whether a stronger actin cable can be observed as a result of increased JNK signaling. As such an analysis of actin cable strength in a JNK gain-of-function genotype has not performed to date, we made time-lapse videos of embryos in which the activity of the JNK pathway is elevated. These experiments were performed on puckered mutants, which provide a well-established and thoroughly characterized JNK gain-of-function genotype (Martin-Blanco et al., 1998; Zeitlinger and Bohmann, 1999; Wang et al., 2003). These embryos lack the Drosophila JNK phosphatase and, therefore, have elevated levels of JNK activity. As shown by the analysis of puc mutant movies, increased JNK signaling results in an apparently more contractile cable (Fig. 2A, Supplementary Video 6), with epidermal bunching toward nodes at the LE. To quantify this effect, we measured the speed with which the actin cable retracts in puc mutant and wild-type embryos after being severed with a laser-directed cut (Fig. 2B, Supplementary Video 7). Quantitative analysis of movies showing the retraction demonstrates that increased JNK signaling correlates with higher tension and contractility of the actin cable during DC (Fig. 2C,D). Taken together, these data confirm that signaling strength correlates with cable thickness and cable tension. The combination of gain- and loss-of-function data strongly suggests that one of the contributions of JNK signaling to the DC process is the assembly of the actin cable in the LE. This developmental effect might involve the transcriptional regulation of genes such as profilin, which we have identified previously as JNK responsive (Jasper et al., 2001). It is likely that JNK signaling has other functions as well, which would account for the pleiotropic phenotype of the respective mutant embryos.

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Figure 2. Increased JNK signaling augments actin cable tension. A: Successive frames (indicated by roman numerals) from a dorsal closure (DC) video of a puc mutant show contractile nodes in the cable (arrowheads in A). B,C: A 5-μm cut at the leading edge (LE; marked by the arrowhead) is made with a ultraviolet laser in similarly staged puc (B) and wild-type (C) embryos. D: The distances between the cut cable edges in B and C are measured and plotted versus time. E: The initial rate of opening (first 20 sec) was averaged over five trials (graphed as average ± standard deviation). This rate of retraction is greater in puc mutants (0.8 ± 0.2 μm/sec for puc compared with 0.5 ± 0.1 μm/sec for wild-type, P < 0.05), providing an indirect comparison of the actin cable tension. Supplementary Videos of A, B, and C are available online (Videos 6,7). Scale bar = 15 μm.

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Next, we wanted to investigate whether the zipping component of DC might also be influenced by JNK signaling. The closure process in the strong loss-of-function conditions described above (jun or slipper null mutants) fails too early for zipping to be reliably observed. Thus, we turned to embryos that are homozygous for the hypomorphic bsk2 allele. bsk2 mutations cause a moderate DC phenotype, with a leading edge actin cable that is stronger than that of an embryo completely deficient for JNK signaling. However, movies of bsk mutant embryos reveal a clear zipping defect. Whereas the canthi in a wild-type embryo move steadily toward the center of the dorsal midline (asterisks in Fig. 1A), such zipping is consistently absent or delayed in bsk2 embryos (arrows in Fig. 1D; see Supplementary Movie 4). Thus, the bsk mutant phenotype indicates that JNK is not only required for actin cable maturation but also for zipping at the canthi.

In the jun or slpr null mutants, the amnioserosa and epithelium rip away from each other causing DC to fail soon after the dorsal rows of epithelial cells have started to stretch (Fig. 1A–C). When the two cell layers have separated, the epithelium falls away ventrally, resulting in the typical final appearance of a “dorsal open” embryo. A similar effect is also seen in hypomorphic bsk mutant embryos, even though it is delayed and occurs after DC has already progressed substantially (Fig. 1D). After the contact between epidermis and amnioserosa is disrupted, the latter contracts and appears to disintegrate (this is only apparent when the amnioserosa remnants stay in the focal plane of observation, such as in Supplementary Videos 3 and 4). The breakdown of the amnioserosa is reminiscent of, and might be mechanistically related to, the removal of this tissue as it normally occurs at the end of DC when contact with the yolk sac is lost. It is unclear at this stage whether such an anoikis-like effect is responsible for the destruction of the amnioserosa after its rupture from the lateral epidermis. However, based on its timing, it seems evident that this is a consequence rather than a cause of failed DC. In summary, at least three phenotypic defects are observed in JNK signaling mutants: lack of proper actin cable formation and contraction, a decrease or absence of zipping, and a loss of attachment between the amnioserosa and the epithelium.

JNK Signaling Regulates βPS Integrin Expression During DC

The marked zipping defect of bsk2 embryos described above might be explained by impaired cell adhesion between the contralateral LE cells at the canthi in the absence of JNK signaling. Potential effectors that could conceivably mediate such changes in cell adhesion are the Drosophila βPS integrin, myospheroid (mys) and the αPS3 integrin scab (scb, Stark et al., 1997). Indeed, mutants for mys have been described previously as dorsal open, a defect that apparently results from a lack of zipping (Brown, 1994; Hutson et al., 2003). Similarly, we find that scb mutants display a specific deficiency in zipping (Fig. 1E and Supplementary Video 5 online), resembling the phenotype of both mys (Brown, 1994; Hutson et al., 2003) and the herein described phenotype of bsk mutants (Fig. 1D). Inspection of similarly staged wild-type, mys, scb, and bsk embryos reveals a marked delay in zipping and canthus progression in the mutants (Fig. 3).

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Figure 3. Zipping is decreased in JNK and integrin mutants. A–H: Phosphotyrosine stains of wild-type (A,E), bsk (B,F), mys (C,G), and scb (D,H) embryos at a similar late stage of dorsal closure (DC) reveals zipping at the canthi in wild-type and a deficiency in zipping in bsk, mys, and scb mutants. Arrows point to areas where the epithelium and amnioserosa are beginning to separate from each other.

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Of interest, by analyzing the genome-wide transcriptional response to JNK signaling, we previously had obtained evidence suggesting that mys was up-regulated by ectopic JNK activation in the embryo, suggesting that mys might be a JNK target gene (Jasper et al., 2001). Thus, based on the phenotypic similarities between bsk, mys, and scb and our genomic evidence, we hypothesized that JNK signaling might stimulate integrin expression to mediate efficient zipping at the canthi. To test this idea, we first examined whether expression of mys and/or scb might be regulated by JNK. To this end, JNK was ectopically activated by expression of Hepact (a constitutively activated form of JNKK) in the embryo using the engrailed-GAL4 driver (Figs. 4, 5). In such embryos, increased expression of Mys protein is detected at the cell surface throughout the embryonic segment expressing Hepact (Fig. 4B,D,F,H, the engrailed domain is marked by coexpression of GFP). In embryos expressing GFP alone (Fig. 4C,E,G), such a cortical stain of Mys protein is not observed (compare Fig. 4E and F, which are at the same cortical level as C and D, respectively). Expression of Hepact in the engrailed domain also induces ectopic expression of scb mRNA by in situ hybridization (Fig. 5A,B). Based on these in situ hybridization and immunofluorescence data, we conclude that JNK signaling can regulate the levels of αPS3 and βPS integrins in epithelial cells.

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Figure 4. JNK signaling regulates Mys expression. A–H: Confocal images of embryos expressing (C,E,G) engrailed-driven GFP alone or (A,B,D,F,H) GFP and Hepact stained for green fluorescent protein (GFP; green), phalloidin (gray), and Mys protein (red). Mys protein is increased at the cell surface of the Hepact domain. C,E,G and D,F,H are at the same plane of section showing the cortical actin cytoskeleton, as stained by phalloidin. I–T: Confocal images of wild-type (I–K, close-ups in O–Q) and bsk (L–N, close-ups in R–T) mutant embryos stained for F-actin (green) and Mys protein (red). Mys protein staining is decreased specifically at the leading edge (LE) of bsk mutant embryos. The phalloidin stain showing the LE and cortical cytoskeleton is at the same plane of section as the Mys stain. Scale bars = 15 μm in N, 5 μm in D,T.

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Figure 5. JNK signaling regulates scb expression. A,B: In situ hybridizations for scb are shown in embryos expressing engrailed-GAL4 alone (A) or engrailed-driven Hepact (B) just after dorsal closure but before cuticle secretion. Ectopic scb RNA is detected in the stripes of Hepact expression. C: Reverse transcriptase-polymerase chain reaction (RT-PCR) -based quantification of scb RNA in dorsal closure (DC) stage embryos expressing either dominant-negative Bsk (BskDN) or Hepact under the control of arm-Gal4 in comparison to flies expressing the driver alone (wt). Simultaneous amplification (duplex PCR) of rp49 was performed as an internal control.

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Next, we examined whether JNK is necessary to direct integrin expression in relevant areas during DC. To do this, we monitored Mys protein expression in wild-type and bsk mutant embryos. Wild-type, DC-stage embryos display Mys protein in the amnioserosa, the LE and at muscle attachment sites (Fig. 4I–K,O–Q). Similarly staged bsk mutants, however, exhibit a decrease in Mys protein specifically in the LE (compare Fig. 4O–Q with R–T). Mys protein levels in the amnioserosa and at muscle attachment sites remain unchanged in bsk mutants (Fig. 4L–N). Therefore, in bsk mutants, JNK signaling appears to be required for inducing expression of Mys only in LE cells, suggesting that JNK controls integrin levels specifically in DC. Similar experiments for Scb could not be performed, as a specific antibody is not available. Semiquantitative reverse transcriptase-polymerase chain reaction (RT-PCR) experiments show, however, that scb mRNA levels are decreased in embryos expressing a dominant-negative form of Bsk and increased when Hepact is expressed (Fig. 5C), indicating that scb is regulated by JNK signaling in a manner that is similar to Mys.

DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

We have shown that, in dorsal closure, a well-established in vivo paradigm for epithelial remodeling, JNK signaling specifies a concerted program of cytoskeletal reorganization and cell adhesion to direct appropriate tissue movements. It is clear that precise balancing of actin cable contraction and cell adhesion/zipping is critical for the ordered movement of cell sheets during DC and presumably other developmental situations as well. The coordinated control of these processes can be expected to determine the shape of developing organs and organisms. As JNK controls both adhesive and cytoskeletal properties, it is well-positioned to control such complex cell behavior.

Our analysis indicates that JNK exerts its control on DC chiefly by transcriptional regulation, as mutants for JNK, JNKK, JNKKK, and Jun appear qualitatively indistinguishable. This conclusion is further supported by a previous report that demonstrated a significant rescue of slpr mutant embryos by expression of an activated form of Jun (Stronach and Perrimon, 2002). Other components, such as Rho family GTPases (Bloor and Kiehart, 2002; Jacinto et al., 2002a), may regulate the cytoplasmic aspects of DC more directly and at the same time induce a transcriptional program that supports the continued process (Glise and Noselli, 1997).

Our studies focused on the function of JNK signaling in the LE where relevant effector molecules such as the integrins and profilin are expressed in response to JNK signaling and where essential aspects of DC undoubtedly take place. At a more complex level, it remains to be determined how spatially restricted signaling interactions specify the morphogenetic movements of DC and how JNK signaling is induced in the first place. In this context, the role of the interface between the amnioserosa and the epidermal LE seems especially intriguing. It has been suggested that signals emanating from the amnioserosa activate the JNK pathway in the LE and trigger the events of DC (Reed et al., 2001; Stronach and Perrimon, 2001; Scuderi and Letsou, 2005). Such signaling interactions between the amnioserosa and the cells of the LE may also underlie an asymmetry of JNK signaling, with higher levels in the leading edge and decreased levels in the amnioserosa, that has been identified as a precondition for DC (Reed et al., 2001). It is possible that the function of integrin expression along the leading edge is to mediate or stabilize the signal exchange between the amnioserosa and the epidermis. Further experiments are required to determine whether integrins might support signaling across the amnioserosa/LE interface directly or by stabilizing adhesion between these two cell sheets.

In addition to its potential function as a source of morphogenetic signals, the amnioserosa also makes mechanical contributions to DC. These contributions, while significantly affecting the rate of closure, are nonetheless dispensable based on laser ablation experiments showing that DC can occur when the structural integrity of the amnioserosa is disrupted (Hutson et al., 2003). It has also been reported that Mys mediates the attachment of the yolk sack and to the amnioserosa (Narasimha and Brown, 2004; Reed et al., 2004). Such a contact was suggested to promote amnioserosa contraction and thereby to contribute to DC. While such a scenario is not inconsistent with our data, it would, by itself, not account for the dorsal open phenotype of JNK mutants. Consistent with studies by Reed et al. (2001) who showed that JNK signaling activity ceases in the amnioserosa at the stage of DC, we find that amnioserosa and yolk sack expression of Mys is not dependent on JNK signaling (Fig. 4 and data not shown) and, thus, probably does not contribute to the JNK mutant phenotype studied here.

Integrin receptors connect the extracellular matrix to the actin cytoskeleton. In addition to the mechanical functions of integrins as mediators of cell adhesion, the proteins have been shown to serve as signal transducing cell surface receptors. It has been speculated that interaction of Myospheroid with its extracellular ligand Pinch, which mediates effects of the integrin in cell adhesion (Clark et al., 2003), might affect JNK signaling negatively (Kadrmas et al., 2004). Of interest, both Mys and Pinch are encoded by JNK responsive genes (Jasper et al., 2001), and a signaling function of integrins upstream of JNK would suggest an attractive feedback mechanism to fine tune the DC process at the level of the cable and zipping components. We did not find, however, any evidence for an effect of mys or scb on the expression of prototypical JNK target genes such as dpp or puc (data not shown), suggesting that such a feedback mechanism, if it were operational, would only make a minor contribution to the execution of DC.

A concerted JNK-mediated regulation of integrin function and actin dynamics at the level of profilin and integrin gene expression, thus, conceivably is important for morphogenetic processes beyond DC. Indeed, JNK as well as Chic and Mys, which we have identified as prototypical JNK effectors in actin polymerization and cell adhesion, respectively, have jointly been implicated in the establishment of planar polarity in the developing eye (Gaengel and Mlodzik, 2003). It will be interesting to investigate whether the underlying regulatory circuitry is conserved among different species and developmental settings.

EXPERIMENTAL PROCEDURES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

Fly Stocks and Genetics

sGMCA transgenic flies express a GFP marker that specifically and ubiquitously labels the cortical actin cytoskeleton (Kiehart et al., 2000). The sGMCA transgene expresses GFP fused to the actin-binding fragment of moesin under the control of the spaghetti squash promoter. Mutant alleles used in this study were jun2 (Kockel et al., 1997), bsk2 (Riesgo-Escovar et al., 1996), slpr921 (Stronach and Perrimon, 2002), pucE69 (Martin-Blanco et al., 1998), mys1, scb2 (obtained from the Bloomington Stock Center). For time-lapse microscopy of mutants, sGMCA was recombined onto each mutant chromosome using standard genetic techniques except for slpr, where slpr/Y;;SGMCA/+ embryos were imaged. Embryos were genotyped using the FM7, act5c-GFP or CyO KrGAL4 UAS-GFP; or TM3 KrGAL4 UAS-GFP balancers (Bloomington). When analyzing fixed samples, a CyO-wg-lacZ balancer was used (kindly provided by S. Bolshakov).

Expression of UAS transgenes was induced using 69B-Gal4, engrailed-GAL4 (Brand and Perrimon, 1993), and armadillo-GAL4 (Bloomington). UAS lines used were UAS-HepAct (Weber et al., 2000) or UAS-GFP (Bloomington).

Germ line clones of jun were created by crossing y w hs-Flp12; sGMCA jun2FRTG13/CyO females to y w hs-Flp12/Y; ovoD1FRTG13/Sco males and heat treating progeny for 2 hr at 37°C once per day during larval development (Chou and Perrimon, 1996). Non-Sco and non-Cyo females that developed from these larvae were crossed to y w hs-Flp12/Y; sGMCA jun2FRTG13/CyO males. Fifty percent of the embryos from this cross exhibited the dorsal open phenotype typical of jun mutants.

Embryo Movies

Embryos were dechorionated in 50% Clorox bleach for 70 sec, washed thoroughly with water, and then mounted in a chamber that contains a gas permeable Teflon window, which allows embryonic development to proceed during image capture (Kiehart et al., 2000). Images were taken on a Zeiss LSM410 or LSM510 or Leica SP2 laser-scanning confocal microscope using a ×40, 1.3 NA oil-immersion objective or ×40, 1.3 NA water immersion or ×40, 1.25 NA oil immersion objective, respectively. Color was added to images using Adobe Photoshop.

Laser Ablation

Laser ablation was performed as described previously (Hutson et al., 2003). Approximately 500 nJ was delivered to the sample per pulse. Measurement of wound sizes was performed using ImageJ software.

Histology

Embryos were immunostained as described previously (Sullivan et al., 2000; Kaltschmidt et al., 2002). Briefly, embryos were dechorionated in 50% bleach for 2 min and fixed in 1:1 heptane:8% formaldehyde for 30 min. Unconjugated phalloidin (Molecular Probes) was added to the fixing solution at a final concentration of 1 μg/ml to stabilize the actin cytoskeleton. Embryos were transferred to double stick tape and devitellineized by hand with a 30-G needle in a Petri dish. Embryos were blocked for several hours in 1% bovine serum albumin and 0.05% Triton X-100 and then incubated overnight with primary antibody diluted in blocking buffer (1:3,000 rabbit anti-β-Gal, Cappel; 1:6 mouse anti-Mys; Developmental Studies Hybridoma Bank). Alexa Fluor 546 phalloidin (Molecular Probes, 1 U/ml final concentration) was added during the secondary antibody incubation for 2 hr at room temperature.

RNA Analysis

Expression analysis by in situ hybridization was performed as described previously (Tautz and Pfeifle, 1989). The compared genotypes were incubated with the same amount of probe and developed side-by-side for the same time period to ensure comparable results.

For semiquantitative PCR, similarly staged embryos were collected and RNA was extracted using Trizol (Invitrogen, Inc.). Equal amounts of RNA were used for first-strand cDNA synthesis using Superscript II Reverse Transcriptase (Invitrogen, Inc.) and oligo (dT)12–18 primer (Intergrated DNA Technologies, Inc.). Duplex polymerase chain reaction was performed using a touchdown protocol for 15 cycles (gradually dropping the annealing temperature each cycle by 1°C from 70 to 56°C) and then an additional 14 cycles at an annealing temperature of 56°C. Primers used for scb were 5′-GTGGCCAAACGAACAGTCAA-3′ and 5′-CAGATCGACACTGTAGCGCA-3′, whereas rp49 primers (for an internal control) were 5′-TCCTACCAGCTTCAAGATGAC-3′ and 5′-CACGTTGTGCACCAGGAACT-3′. PCR products were analyzed on a 2% agarose gel.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

We thank B. Stronach, M. Mlodzik, W. Li, the Bloomington Stock Center, and the Developmental Studies Hybridoma Bank for gifts of fly stocks and antibodies. Special thanks to R. Montague for making videos for quantitative analysis. Thanks also to S. Hong, C. Ovitt, C. Wang, R. Ambrosini, C. Sommers, and A. Boury for experimental help and critical comments on the manuscript. J.G.H. is supported by the U of R MD/PhD Program. D.P.K. is supported by NIH and the Department of Defense/Air Force Office of Scientific Research, Medical Free Electron Laser.

REFERENCES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
  9. Supporting Information

The Supplementary Material referred to in this article can be found at http://www.interscience.wiley.com/jpages/1058-8388/suppmat

FilenameFormatSizeDescription
jws-dvdy.20649.fig1.tif4644KSupporting Information file jws-dvdy.20649.fig1.tif
jws-dvdy.20649.vid1.mov4086KSupporting Information file jws-dvdy.20649.vid1.mov
jws-dvdy.20649.vid2.mov5099KSupporting Information file jws-dvdy.20649.vid2.mov
jws-dvdy.20649.vid3.mov3297KSupporting Information file jws-dvdy.20649.vid3.mov
jws-dvdy.20649.vid4.mov3034KSupporting Information file jws-dvdy.20649.vid4.mov
jws-dvdy.20649.vid5.mov9136KSupporting Information file jws-dvdy.20649.vid5.mov
jws-dvdy.20649.vid6.mov4838KSupporting Information file jws-dvdy.20649.vid6.mov
jws-dvdy.20649.vid7.mov1053KSupporting Information file jws-dvdy.20649.vid7.mov

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