Hedgehog Signaling Is a Principal Inducer of Myosin-II-Driven Cell Ingression in Drosophila Epithelia

Developmental Cell

Article Hedgehog Signaling Is a Principal Inducer of Myosin-II-Driven Cell Ingression in Drosophila Epithelia Douglas Corrigall,1 Rhian F. Walther,1 Lilia Rodriguez,1 Pierre Fichelson,1 and Franck Pichaud1,* 1MRC Laboratory for Molecular Cell Biology and Cell Biology Unit, Department of Anatomy and Developmental Biology, University College London, Gower Street, WC1E 6BT, London, United Kingdom *Correspondence: [email protected] DOI 10.1016/j.devcel.2007.09.015

SUMMARY

Cell constriction promotes epithelial sheet invagination during embryogenesis across phyla. However, how this cell response is linked to global patterning information during organogenesis remains unclear. To address this issue, we have used the Drosophila eye and studied the formation of the morphogenetic furrow (MF), which is characterized by cells undergoing a synchronous apical constriction and apicobasal contraction. We show that this cell response relies on microtubules and F-actin enrichment within the apical domain of the constricting cell as well as on the activation of nonmuscle myosin. In the MF, Hedgehog (Hh) signaling is required to promote cell constriction downstream of cubitus interruptus (ci), and, in this context, Ci155 functions redundantly with mad, the main effector of dpp/BMP signaling. Furthermore, ectopically activating Hh signaling in fly epithelia reveals a direct relationship between the duration of exposure to this signaling pathway, the accumulation of activated Myosin II, and the degree of tissue invagination. INTRODUCTION Organogenesis requires the precise regulation of cell proliferation, movement, and apoptosis to achieve proper organ shape and size. In addition, the folding of epithelial cell sheets plays a crucial role in shaping organs such as the heart, lung, and kidney. In many organisms, epithelial-cell sheet invagination is promoted when a few epithelial cells adopt a bottle- or wedge-like shape resulting from a striking constriction of their apical domain. Such cellshape changes create a local mechanical stress within the epithelium that is instrumental for promoting tissue invagination (Kimberly and Hardin, 1998). This is likely to be a conserved feature in tissue patterning because the formation of bottle-shaped cells associated with tissue invagination has been identified in the sea urchin during

gastrulation (Davidson et al., 1995), during neural tube closure in the vertebrate notochord (Schroeder, 1970), and in the developing fly embryo (Kiehart et al., 1990), to name but a few examples. Apical cell constriction is crucial for mesoderm invagination during Drosophila development, and, in the past decade, key molecular players involved in this process have been identified. In the fly embryo, cell constriction is characterized by an accumulation of apical F-actin and activation of hexameric, nonmuscle MyoII through the RhoA-Rok signaling pathway (Dawes-Hoang et al., 2005; Hacker and Perrimon, 1998; Nikolaidou and Barrett, 2004; Seher et al., 2006). Cell constriction is achieved through the assembly of MyoII into bipolar filaments and the assembly of short bundles of unbranched F-actin that act as a substrate for the motile activity of MyoII. Rok appears to be the principle kinase that activates MyoII via phosphorylation of a conserved serine at position 19 of the myosin regulatory light chain (MRLC) (encoded by spaghetti squash [sqh]) (Amano et al., 1996). Activated MyoII is then thought to be involved in pulling the adherens junctions (AJs) toward the apical pole of the cell, thereby ‘‘eating-up’’ the apical surface (Dawes-Hoang et al., 2005; Kolsch et al., 2007). In addition, a recent report revealed that transient AJ disassembly also takes place during cell constriction (Kolsch et al., 2007). In the fly embryo, ventral furrow formation involves the transcription factor Twist upstream of the G protein Concertina pathway (Parks and Wieschaus, 1991; Seher et al., 2006) as well as the recently identified transmembrane protein T48 (Kolsch et al., 2007). In this context, the putative secreted protein Folded gastrulation induces the cytoskeletal changes required to promote tissue invagination (Costa et al., 1994; Morize et al., 1998). However, knowledge about the upstream signaling pathways that provide the patterning information that governs cell constriction and eventually cell ingression during organogenesis remains scant. A situation strongly reminiscent of cell constriction is cell cytokinesis. Specifically, the assembly of the contractile ring necessary for the function of the cleavage furrow and cell division also requires the assembly of parallel F-actin and motile forces provided by MyoII (Dean et al., 2005; Hickson et al., 2006). Additionally, the MyoII-driven motile force required to achieve cell cleavage involves the RhoA-Rok signaling pathway (Glotzer, 2001). In this

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context, the microtubule-associated and F-actin-nucleating formin Diaphanous (Dia) is required for stabilizing MyoII at the cleavage furrow. This is achieved through the actin-nucleating activity of the formin homology (FH) 2 domain present in Dia and is presumably combined with the ability of this protein to directly interact with Profilin, another F-actin effector (Chang et al., 1997). However, a role for dia in cell constriction remains to be examined. To gain insight into how cell constriction and tissue invagination are regulated during development, we turned to the genetically amenable Drosophila eye imaginal disc. Patterning of this columnar pseudostratified epithelium depends on the formation and movement of the morphogenetic furrow (MF) along the anterior-posterior axis. The MF is characterized by cells undergoing a synchronous apical constriction and apicobasal contraction together with a cell-cycle arrest in G1 (Ready et al., 1976; Wolff and Ready, 1991). Hh and Dpp signaling are instrumental for propagating the MF; Hh is produced by the developing neurons and then diffuses more anteriorly (Figure 1A). The two signaling pathways rely on transcriptional regulation of genes downstream of Ci and Mad, respectively (Lum and Beachy, 2004). We show that the cell constriction and mild ingression observed in the MF are transcriptionally controlled downstream of Hh. This signaling pathway requires the small GTPase RhoA and its effector, Rok, to control the activation of nonmuscle MyoII, whereas Dia, coupled with the regulated activity of Cofilin and Profilin, is required for proper F-actin enrichment within the apical cortex of the constricting cells. Our data are consistent with a model in which the stabilization of apical MTs in the MF is coupled to the assembly of parallel F-actin within the apical cell cortex and in which the motile force necessary for apical constriction is provided by MyoII. Importantly, our work suggests a dose-dependent relationship between the level of phosphorylated MyoII within the apical membrane and the types of structures produced in a columnar epithelium. Short-term exposure to Hh signaling leads to cell constriction and mild ingression, whereas longer exposure to this pathway is accompanied by higher levels of activated MyoII and leads to major tissue invagination. RESULTS Cell Constriction in the Morphogenetic Furrow Requires RhoA/Rok and MyoII Cells in the MF are characterized by a striking apical cell constriction (Figures 1A–1C) and apico-basal contraction (Figures 1D–1F), which can be viewed as a mild case of tissue invagination (Ready et al., 1976; Wolff and Ready, 1991). This is accompanied by an increase in F-actin and E-cadherin (encoded by shotgun [shg]) within the apical cortex of the constricting cells (Figures 1A and 1H). Interestingly, we also observed a substantial stabilization of the apical MT network (Figures 1E and 1F) in the constricting cells in a region spanning the apical domain and septate junctions (SJs), which are located basal to the AJs. These cytoskeletal changes are accompanied by a dra-

matic reduction of the apical cell surface area (Figures 1A–1C) also apparent at the level of the basal SJs, as seen by labeling with an antibody against Discs-large (Dlg) (not shown). Whereas the MF is characterized by an acute decrease in the apical cell surface of over 10 cell diameters, the whole anterior compartment of the eye disc is characterized by a graded decrease of the apical cell surfaces along the anterior-posterior axis (Figures 1A and 1B); smaller apical surfaces are measured closer to the MF. This suggests that the MF response is distributed over the entire anterior compartment of the developing eye. In addition, optical sagittal sections (Figure 1D) revealed that, in the anterior compartment, cells become increasingly taller along the anterior-posterior axis; cells in the anterior-most part of the epithelium are more cuboidal than the columnar cells close to the MF. Consistent with the hypothesis that MyoII could be required for cell constriction in the MF, we observed MRLC enrichment at the apical cortex of the constricting cells in the MF (Figure 1G), where it is activated through phosphorylation of Ser19 (Figures 1H–1K). We next used the FLP/FRT system (Xu and Rubin, 1993) in the developing eye to examine the effects of removing the function of genes previously shown to activate and modulate MyoII activity. Removing rok function in the MF caused a marked decrease in apical F-actin quantity and a statistically significant (p < 0.05) impairment in apical cell constriction (WT-MF cell surface/rok cell surface: 0.539) (Figures 2A and 2B; Tables S1 and S2, see the Supplemental Data available with this article online for quantification) and apico-basal contraction (Figures 2C and 2D). This was accompanied by a statistically significant (p < 0.05) decrease in Ser19 MRLC phosphorylation (Figures 2E–2H; Table S3 for quantification). Nevertheless, we could detect apical Myosin Heavy Chain (Zipper [Zip]) in these mutant cells (Figures 2A and 2D and data not shown). Importantly, epithelial cell polarity was maintained in these mutant cells (not shown), indicating that the impaired MF cell response was not caused by a loss of polarity. Although Rok function is key for cell constriction, expressing an activated form of this kinase (RokCAT) (Winter et al., 2001) in clones was not sufficient to promote cell constriction in the anterior compartment of the eye disc (Figures S1A–S1C). We next examined mutant cells deficient for MRLC (sqh). We were only able to recover small mutant clones by using a hypomorphic allele of MRLC, probably because MRLC is required for normal cytokinesis (Edwards and Kiehart, 1996). In these mutant clones, we observed a failure in the MF cell response in that both constriction and apicobasal contraction failed to occur (Figures 2I–2L and data not shown). Similarly, cell constriction and apico-basal cell contraction failed to occur when a dominant-negative version of MRLC lacking the F-actin-binding domain (DawesHoang et al., 2005) was expressed in clones (not shown). Interestingly, removal of the Myosin phosphatase-binding subunit (mbs) in mosaic eye discs led to groove formation in the case of clones located in the MF (Figures 2M–2P0 ). This finding suggests that different levels of MRLC phosphorylation might promote different degrees of tissue

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Figure 1. Epithelial Morphogenesis and Drosophila Eye Organogenesis All of the tissue samples are orientated along the anterior-posterior axis. The MF is marked with a solid, white line. (A) Apical view of a developing eye labeled for the AJ marker DECadherin:GFP. The anterior compartment is indicated by a yellow line, whereas the proneuronal compartment and MF are indicated by turquoise and blue lines, respectively. The posterior compartment is indicated by a purple line. (B) Representation of the apical cell surface measured along the anterior-posterior axis of the developing eye (n = 5 discs). A red line marks the MF. (C) Schematic representation of a columnar epithelial cell found in the anterior compartment ahead of the MF (left) and a constricting cell found in the middle of the MF (right). (D) Sagittal section of a developing eye labeled for F-actin (red) and a marker of the SJs, Dlg (green). ppm, peripodial membrane. (E) High magnification of the eye primordium centered on the MF. The F-actin is labeled in red, whereas the microtubules (MTs) are labeled with a-tubulin (green). (F) Same as (E), but showing the MTs only. (G) A fusion protein between MLRC and GFP is used to report MyoII localization (green). PATJ (blue) labels the apical domain, and Dlg (red) labels the SJs. (H–K) Phosphorylation at Ser19 of MLRC (green) in the MF. Arm marks the AJs (blue); phalloidin labels F-actin (red).

invagination, with higher levels associated with tube formation. Importantly, high levels of expression of an activated form of MRLC (SqhE20E21) in the anterior compart-

ment of the developing eye disc proved to be sufficient for inducing reproducible cases of minor cell constriction accompanied with tissue ingression (Figures 2Q–2T).

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Figure 2. Patterning of the Eye Imaginal Disc: MF and MyoII-Driven Cell Constriction In all panels, the MF is indicated by a white line, and the clones, marked by the absence of blue (b-galactosidase or GFP staining), are circled with a dashed, white line. (A and B) Cells mutant for the null allele rok2 showing a strong reduction in apical F-actin staining (red) correlated with a failure to properly achieve cell constriction. The dashed lines labeled (C) and (D) mark the position corresponding to the sagittal sections seen in (C) and (D), respectively. (C and D) Sagittal sections showing a failure of the (D) rok2 mutant cells to undergo apicobasal contraction, compared to the (C) wild-type tissue shown in the same preparation. (E–H) Cells mutant for the null allele rok2. Factin staining is shown in red, and PhosphoSer19 MRLC staining is shown in green. The cell nuclei are labeled with DAPI (white) in (H). (I–L) Cells mutant for MRLC (sqh1). F-actin staining is shown in red, and Dlg staining is shown in green. (M–P0 ) Eye disc presenting a clone mutant for mbs791. (M)–(P) show an optical section at the level of the apical membrane of the constricting cells in the MF, and the position of the mutant clone is indicated with the dashed line. (M0 – P0 ) show a basal view of the same region of the eye disc. This reveals that the apical membranes of the mbs791 mutant cells are located at the level of the basal nuclei (F-actin staining is shown in red; the AJ marker Arm staining is shown in green). (Q–T) Activated MRLC (SqhE20E21) induces mild cell ingression (lack of green staining). F-actin staining is shown in red; Nuclei are stained blue. An arrowhead points to the ectopic cell ingression. (U–X) Eye disc mutant for RhoA72M1 induced by using the Minute technique. MRLC phosphorylation at Ser19 is shown in green.

Finally, in the absence of RhoA function, we observed a dramatic loss of cell constriction and apicobasal contraction in the MF, associated with weak F-actin staining and a diffuse pattern of MRLC phosphorylation at Ser19 when compared to wild-type (Figures 2U–2X). Taken together, our findings demonstrate that cell constriction and ingression in the MF depend upon the action of RhoA, Rok, and MyoII. Our observation of residual phosphorylation of MRLC at Ser19 in rok mutant cells (Table

S3) confirms a previous report (Lee and Treisman, 2004) and suggests that another kinase acts in parallel with Rok to phosphorylate this residue. The Formin Dia Is Required for Apical Cell Constriction One major effect of removing RhoA and rok function in the MF is a significant reduction in F-actin accumulation in the apical domain of the cell (Figures 2B and 2V; Table S2 for

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quantification). Previous work on the regulation of cytokinesis has revealed that RhoA acts upstream of the formin Dia (Dean et al., 2005; Hickson et al., 2006). In addition, together with Dia, both profilin (encoded by chickadee [chic]) and ADF/cofilin (encoded by twin star [tsr]) have been shown to be important for cytokinesis through promoting F-actin reorganization at the contractile ring and thus providing a substrate for MyoII-mediated motility (Ishizaki et al., 2001; Palazzo et al., 2001). This prompted us to test whether dia could function in apical constriction. First, we showed that Dia is localized in the apical domain of constricting cells (Figures 3A–3C). We then showed that the removal of dia completely inhibited apical constriction in the MF (Figures 3D–3G) and largely prevented F-actin enrichment in the mutant cells (Figures 3E and 3G). Together, these observations suggest that cell constriction is dependent upon Dia-driven enrichment of actin filaments in the apical domain of these cells. However, ectopic expression of a constitutively activated form of Dia, Dia-CA (Somogyi and Rorth, 2004), failed to trigger cell constriction or MT stabilization in the corresponding expressing cells in the eye and wing imaginal discs (not shown). We then examined clones mutant for profilin. As previously reported in the developing eye (Lee and Treisman, 2004), a clear loss of apical F-actin is observed in the corresponding mutant cells (Figures 3H–3K). However, we also noted that this was accompanied by an alteration of cell shape and a substantial loss of apico-basal polarity, including a strong decrease in the SJ marker Dlg (Figure 3J). We next assayed a role for ADF/cofilin and, as previously reported (Lee and Treisman, 2004), we noted a strong statistically significant (p < 0.05) increase in cortical F-actin in the corresponding mutant cells (Figures 3L–3O; Tables S1 and S2). The corresponding apical cell surface areas were significantly (p < 0.05) enlarged (Tables S1 and S2). A similar situation was obtained when removing the cofilin phosphatase slingshot (ssh) in clones (Figures 3P–3S; Tables S1 and S2). The increase in cell surface area measured in cofilin and ssh mutant cells in the MF is likely to be the result of these cells being filled with a large excess of F-actin that spreads basally toward the SJs (not shown). This set of data indicates that Dia, Profilin, and Cofilin act in concert to promote apical Factin enrichment in parallel with MyoII, which is needed for cell constriction. Our data are compatible with the possibility that both MyoII and dia act downstream of rok and RhoA, a situation strongly reminiscent of that observed during cytokinesis (Dean et al., 2005). Cell Constriction and Actin Effectors A recent study has revealed a role for the nonreceptor and actin-binding Abelson tyrosine kinase (Abl) and the actin effector Enabled (Ena) in promoting cell constriction in the fly mesoderm (Fox and Peifer, 2007). However, we could not detect any defect in cell constriction in the MF when inducing mutant clones for either Abl or ena by using null alleles for these two genes (Abl4 and ena23, respectively) (Figures 3T–3W and 3X–3A0 ). This might be due to

some redundancy in this cell response, and, in that respect, it is interesting to note that in the mesoderm, cells undergoing apical constriction could be detected in the absence of Abl function (Fox and Peifer, 2007). Similarly, we failed to detect defects in apical constriction in the MF when generating loss-of-function mutant cells for DWave (using a null allele, scarD37) (Figures 3B0 –3E0 ) and Wasp (using a combination of independent alleles, wsp1 and wsp3; data not shown), both key effectors of F-actin and regulators of the Arp2/3-actin-nucleating complex (Machesky et al., 1994; Miki and Takenawa, 2003; Nakagawa et al., 2001). It is also interesting to note that, despite our efforts with various allelic combinations and loss-offunction mutant clones, we failed to detect defects in cell constriction in mutant cells for Rho-GEF2 (Figures S1E–S1G), a conserved Guanosine Exchange Factor (GEF) responsible for loading GTP onto Rho1/A. When generating whole mutant eye discs for Rho-GEF2 we could readily detect defects in the gross morphology of the disc, including the folding of the corresponding disc onto itself (data not shown). These data suggest that, in the MF, another GEF might function in parallel with RhoGEF2, or that Rho-GEF2 is not involved in cell constriction in the MF. Consistent with an important role for the AJs during cell constriction, mutant clones for arm induced in the MF led to a failure in this cell response (Figures S1H–S1K). Hh Signaling Is a Principal Inducer of Apical Cell Constriction We next investigated the potential upstream regulator(s) responsible for orchestrating the MF cell response. It has previously been reported that both the Dpp and Hh pathways provide spatial and temporal patterning information in the eye imaginal disc (Chanut and Heberlein, 1997; Borod and Heberlein, 1998). Hh and Dpp signaling are involved in MF induction and propagation, but it is not clear if both of these pathways are required for cell constriction in the MF. To address this issue, we induced loss-of-function clones in the developing eye disc for mothers against dpp (mad; vertebrate smad5/7) (Figures 4A–4D; Figure S1D), the Hh coreceptor smoothened (smo) (Figures 4E–4H and 4M–4P), or both mad and smo (Figures 4I–4L, 4Q–4T, and 4U–4X). As previously reported in the wing disc (Gibson and Perrimon, 2005; Shen and Dahmann, 2005), removing mad function in the eye by using a null allele led to clones producing cysts that sometimes could be recovered in the basal part of the epithelium (Figure S1D). However, we could also recover a number of clones in the MF that were relatively small and did not produce cysts. In these clones, we did not detect significant changes (p > 0.05; Table S1) in cell constriction (cell surfaces WT-MF/mad: 0.82), F-actin accumulation, or MRLC-Ser19 phosphorylation at the apical cortex (Figures 4A–4C; Tables S1–S3). The fraction of mad mutant clones that we were able to recover in the MF did not affect the characteristic apico-basal contraction of the corresponding cells (Figure 4D). Conversely, in smo mutant clones, we consistently observed a strong and statistically

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Figure 3. Cell Constriction Depends on the Activity of Dia, Profilin, and Cofilin (A–C) Sagittal section of an eye imaginal disc. Dia (blue) colocalizes with MLRC:GFP (green) and the AJ marker Arm (red). (D–G) Cells mutant for dia5. F-actin staining is shown in red, and Dlg marks the SJs (green). (H–K) Cells mutant for profilin (chic221). (L–O) Cells mutant for cofilin (tsr1). (P–S) Cells mutant for ssh221. (T–W) Cells mutant for Abl4. F-actin is labeled in red, whereas the AJ marker Arm is in green. (X–A0 ) Cells mutant for ena23. F-actin is labeled in red, whereas the neuronal marker Elav is in green. (B0 –E0 ) Cells mutant for SCARD37. F-actin is labeled in red, whereas the SJ marker Dlg is in green.

significant (p < 0.05) impairment in apical cell constriction in the MF (cell surfaces WT-MF/smo: 0.43; Figures 4E–4H and Table S1), which was accompanied by a dramatic

decrease in F-actin accumulation (Figure 4F; Table S2) and a marked decrease in b-catenin (Armadillo [Arm]) accumulation at the AJs (data not shown). In addition, we

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Figure 4. Hh Signaling Regulates MyoIIDriven Cell Constriction (A–C) Cells mutant for mad12. F-actin is labeled in red. (D) Sagittal section across a mad12 mutant clone showing that cells still achieve proper apicobasal contraction (marked with a white line). (E–H) Cells mutant for smoD16 exhibiting a failure in apical F-actin enrichment (red). Dlg marks the SJs (green). (I–L) Double mutant clone for mad1-2 and smo3 exhibiting a failure in apical F-actin enrichment (red). Dlg marks the SJs (green). (M–P) smoD16 mutant cells showing a total loss of MRLC phosphorylation at Ser19 (green). (Q–T) Double smo3, mad1-2 mutant cells exhibiting failure to achieve apical constriction (F-actin, red) and a total loss of MRLC phosphorylation at Ser19 (green). (U–X) Loss of Dia staining (green) at the apical cortex in the MF in the absence of smo3 and mad1-2 function. F-actin is labeled in red, and the mutant cells are indicated with a white asterisk.

measured a significant decrease (p < 0.05) in the levels of MRLC phosphorylation at Ser19 at the apical cortex of these mutant cells (Figure 4Q; Table S3), implying that Hh signaling plays a major role in inducing cell constriction and apico-basal contraction in the MF. However, by using Dlg to label the SJs, we could still observe a remnant of the MF (Figure 4G), suggesting that Dpp and Hh might act in a partially redundant manner in the control of the MF cell response. To test this hypothesis, we removed both of these two signaling pathways in clones mutant for both mad and smo. The mutant cells completely failed to undergo apical constriction (cell surfaces WT-MF/mad, smo: 0.30; Figures 4I–4L and Table S1) and apico-basal contraction. Furthermore, there was no F-actin accumulation at the apical cortex (Table S2) or MRLC phosphoryla-

tion at Ser19 (Figures 4Q–4T; Table S3). In these mutant cells, we also observed a substantial decrease in apical Dia staining (Figures 4U–4X). Nonetheless, cell polarity was preserved, as assayed by using marginal zone, AJ, and SJ markers (Figure 4K and data not shown). Apical Cell Constriction in the MF Is Transcriptionally Regulated Downstream of Hh To further test the function of Hh signaling in cell constriction, we removed the ci locus (i.e., both Ci155-transcriptional activator and Ci75-transcriptional repressor), the zinc finger transcription factor responsible for mediating Hh signaling downstream of Smo (for a review, see [Lum and Beachy, 2004]). In ci mutant clones induced in the anterior compartment of the eye imaginal disc, we

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Figure 5. Hh Signaling Transcriptionally Regulates Cell Constriction and Ingression (A–D) Cells mutant for ci showing ectopic apical constriction immediately anterior to the endogenous MF. F-actin (red) and Dlg labeling the SJs (green). (E) Sagittal section of a cullin116E clone. (F–H) Double mutant clone for the transcription factor ci and mad1-2 fail to show ectopic apical constriction. (I–M) Sagittal section across a clone mutant for ci (loss of green staining, dashed lines). (I) A phospho-specific antibody labels P-Mad (blue); (K) F-actin staining is shown in red, and the (I) cell nuclei are labeled with DAPI (white). (N–R) (N) Sagittal section across an eye disc expressing spastin:GFP. (O) P-Ser19 staining (blue). (P) MT staining in red. (R) Nuclei are stained with DAPI (white).

consistently observed ectopic MF-like cell responses, including apical MT stabilization and MyoII activation (Figures 5A–5D and not shown). Similarly, we prevented Ci75-transcriptional repressor formation by removing cullin1, a ubiquitin-ligase required to generate the transcriptional repressor Ci75 from the transcriptional activator Ci155 (Ou et al., 2002). cullin1 loss-of-function clones led to ectopic MF cell responses in the anterior compartment immediately ahead of the MF (Figures 5E). These experiments suggest that alleviating Ci75-mediated transcriptional repression is sufficient to induce cell constriction. Surprisingly, while mad is not absolutely required for a cell to constrict in the MF, loss of mad function in ci mutant cells (in double ci and mad mutant cells) led to a lack of ectopic cell constriction in the corresponding cells (Figures 5F–5H). This indicates that, in the absence of Ci155-mediated transcriptional activation, ectopic cell constriction requires Dpp signaling to proceed. Although, all of the cells located in the anterior compartment of the eye disc receive the Dpp signal, as reported by a phospho-specific antibody raised against active P-Mad (Figures 5I–5M), we did not observe increased P-Mad staining in ci mutant clones. Altogether, this suggests that mad function is required for cell constriction to occur in ci mutant cells, whereas in the MF, Ci-155 and Mad might act redundantly to promote this cell response. In ci mutant cells and in the endogenous MF, cell constriction is accompanied by MT stabilization within the apical cell domain. To assay their contribution during cell

constriction, we used the MT-severing factor Spastin (Sherwood et al., 2004) to depolymerize MTs in vivo (Figures 5N–5R). When expressed in the MF (Figure 5N), Spastin led to a near total depletion of the MT network, accompanied by a loss of apical F-actin accumulation and a loss of Ser19-P MyoII staining. In addition, the basally located nuclei normally found in the MF appeared randomly distributed and had a tendency to be close to the apical cell surfaces (Figure 5Q). Hh Signaling Induces a Range of Tissue Invagination We next tested whether Hh signaling was sufficient to trigger cell constriction in fly epithelia. To this end, we ectopically activated the Hh signaling pathway in the eye, antenna, and wing imaginal discs by generating mutant clones in which the inhibitory coreceptor patched (ptc) was removed (Hooper and Scott, 1989). In ptc mutant clones, Hh signaling is constitutively activated at its maximal strength. Interestingly, these clones led to a range of responses, depending on the levels of activation of MyoII measured through quantifying the Ser19-P MRLC staining (Table S3). In the case of ptc clones dissected 24 hr after clone induction, we invariably observed MF-like responses (Figures 6A–6D; Tables S1 and S2) together with levels of Ser19-P MRLC comparable to those measured in the endogenous MF cells (p > 0.005; Table S3). This was accompanied by a pronounced bundling of the apical MTs (Figures 6E–6H) and a striking apical

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Figure 6. Hh Signaling Promotes MyoIIDriven Tissue Ingression (A–D) Sagittal sections showing a mild case of cell ingression for ptcS2 mutant cells in the eye disc. F-actin staining is shown in red, and Ser19-P-MRLC staining is shown in green. The endogenous MF is indicated by a white line in (B) and (C) and by a red line in (D). (E–H) ptcS2 mutant clones induced in the anterior compartment of the eye showing F-actin (red) and MT stabilization (green). (I–L) Sagittal section of a ptcS2 mutant clone induced in the wing disc and leading to tissue invagination. F-actin staining is shown in red, and Ser19-P-MRLC staining is shown in green. (M) Sagittal section of a ptcS2 mutant clone induced in the eye disc and leading to groove formation. F-actin staining is shown in red, and Dlg staining is shown in blue. (N) Schematic representation of the endogenous MF cell response. Blue labels the dpp-expressing cells, whereas orange labels the Hhproducing cells. (O) Schematic representation of the situation described in (L) upon removal of patched (ptcS2). This leads in the wing to alleviation of Ci75-mediated transcriptional repression, and it allows for Ci155-mediated transcriptional activation as well as the induction of dpp expression (Methot and Basler, 1999).

enrichment of F-actin (Figures 6F; Table S2). This was not limited to the eye imaginal disc because removing ptc function in the antennal or wing discs also resulted in ectopic activation of MyoII and enrichment of apical F-actin and MT stabilization, leading to a striking cell constriction in the affected tissue (Figures 6I–6L and not shown). Interestingly, when ptc mutant clones were dissected 48–72 hr after their induction, we primarily observed an ingression of the ptc mutant cells, with the cells dropping down within the columnar epithelium, without loss of polarity or loss of adhesion. This was the case in all imaginal discs, including the wing (Figures 6I–6L) and the eye (Figure 6M). As expected, the ptc mutant cells showed an increase in apical F-actin and MRLC phosphorylation at Ser19 (Figure 6J, 6K, and 6M). Quantification of Ser19 phosphorylation of MRLC in the eye revealed that cells located at the bottom of such ptc-induced tubes exhibit a significantly higher level (p < 0.05) of activated MyoII when compared to the endogenous MF cells (Table S3). These observations suggest that Hh signaling can induce cell ingression and groove formation in a MyoII-dependent manner (Figures 6N–6O; Table S3) provided there are a minimum number of Hh signaling cells and adequate levels of activated MyoII.

DISCUSSION Hh Signaling and Tissue Patterning The Hh and Dpp families of ligands are of crucial importance for patterning a wide variety of tissues across species. Our work on Hh signaling and that of others on Dpp (Gibson and Perrimon, 2005; Shen and Dahmann, 2005) suggest that these major signaling pathways also function in modulating MTs, F-actin nucleation, and myosin activity during organogenesis. When removing ci (both Ci155-act and Ci75-rep), we consistently observed ectopic MF-like responses in the eye imaginal disc, even when generating relatively large patches of mutant cells. The failure to induce apical constriction in double ci, mad clones demonstrates that transcription downstream of Dpp signaling is required for ectopic cell constriction to proceed in the absence of Ci155. As mad function is dispensable in the MF, this also suggests that, in this context, Ci155 and Mad might function redundantly. We found that a number of mad mutant clones form cysts in the eye disc and, as previously reported (Gibson and Perrimon, 2005; Shen and Dahmann, 2005), this is accompanied by defects in apical MT organization. It is not clear why some of the mad mutant clones lead to cyst

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formation while others remain within the epithelium, and it is possible that the timing of mutant-clone induction is a key parameter. Interestingly, cells in the MF present the highest levels of P-Mad, and this correlates with our observation that, in these cells, the apical MTs appear more bundled and abundant. Linking MT and F-Actin during Cell Constriction The Formin-Homology (FH) protein family has been implicated in a number of actin- and MT-dependent cellular processes, including the alignment of the MTs and F-actin during stress-fiber formation (Tsuji et al., 2002), cell polarity, and cytokinesis (Wasserman, 1998). In the fly embryo, dia loss of function leads to defects in pseudocleavagefurrow invagination during cellularization (Afshar et al., 2000) and contractile-ring morphogensis (Giansanti et al., 1998). Consistent with a role for dia during cytokinesis, we observed that a high proportion of dia mutant cells were located at the apical surface of the epithelium and arrested in telophase. Nevertheless, we could recover small dia mutant clones in the MF that retained their overall apico-basal polarity, thus enabling us to reveal a function for this factor for F-actin enrichment during cell constriction in the MF. However, we were not able to induce ectopic cell constriction by using an activated form of dia (Somogyi and Rorth, 2004). FH proteins such as Dia contain a central proline-rich FH1 domain that, in yeast, has been shown to mediate a direct interaction with the actin effector Profilin (Chang et al., 1997), whereas the amino-terminal part of several FH proteins is able to interact with the GTP-bound RhoGTPases such as Rho1 (Watanabe et al., 1997, 1999). Our data suggest that a RhoA-Dia-Profilin biochemical module might be evolutionarily conserved (Chang et al., 1997). Interestingly, mdia has been shown to be transported on MT and act as a MT (+) end stabilizer in 3T3 fibroblasts (Palazzo et al., 2001). Whereas MTs remain largely unaffected in the absence of dia in the MF (not shown), depolymerizing MTs (Sherwood et al., 2004) led to the absence of cell constriction in the MF, accompanied by a loss of apical F-actin and Ser19-P MyoII staining. In this context, we favor a model in which dia is involved in maintaining F-actin at the apical cell cortex in the MF, but is not required for the apparent tight bundling and stabilization of the apical MTs in these cells. Finally, it is interesting to note that, in the fly syncytial blastoderm embryo, F-actin and MyoII are transported toward the MT (+) end (Foe et al., 2000). The failure in apical F-actin and activated MyoII enrichment when MTs are depolymerized in the MF is consistent with a role for MTs in targeting MyoII to the apical cell surface. MyoII Activation and Cell Constriction In the MF, we note that loss of rok function does not lead to a total loss of phosphorylation of MRLC at the conserved Ser19. This is consistent with a previous report in the developing eye disc (Lee and Treisman, 2004) and suggests that other kinases act in parallel with Rok in the MF. Potential candidates are the P21-activated kinases (PAK),

encoded by three loci in Drosophila (PAK1/PAK, DPAK2/ mbt, and PAK3). We used the null allele mbtP1 as well as various PAK1 alleles (including PAK16, which encodes a truncated protein) to examine the role of these kinases in apical cell constriction in the MF, and, in both cases, we failed to detect defects in apical cell constriction (not shown). In addition, expressing the auto-inhibitory domain (PAK-AID) found in the group-I PAKs (including PAK1 and PAK3) in clones, and thus interfering with these two kinases’ function (Conder et al., 2004), did not lead to loss of cell constriction in the MF (not shown). It is possible that residual DPAK2/mbt activity could compensate for the lack of PAK1 and PAK3 function, and further work will be required to analyze the effects of removing all three of these loci in the MF. Another kinase potentially responsible for phosphorylating MRLC in the MF is myosin light chain kinase (MLCK), a large, complex locus for which lossof-function mutations have not been identified. MyoII Activation and Tissue Invagination In ptc mutant clones, we are ectopically inducing maximum levels of Hh signaling. In this context, MF-like ingression (or indentation) is associated with lower levels of Ser19-P-MyoII than those we measured in cases of groove formation. Interestingly, the situation in the endogenous MF involves a very transient stimulation of the Hh pathway due to the activity of the ubiquitin ligases Cullin1 and Cullin3, which process Ci155 in the anterior and posterior edge of the MF, respectively (Ou et al., 2002). Our data are consistent with a model in which, upon Hh signaling, activated MyoII accumulates at the apical cortex of the constricting cell over time. The degree of activated MyoII accumulation is linked to the duration of Hh signaling perceived by the corresponding constricting cells and appears to be the main parameter involved in groove formation in columnar epithelia. It is likely that the Hh-induced MyoII-driven pathway we report here also plays a role in tissue patterning in higher organisms. For example, cell constriction and apico-basal contraction occur during the invagination of the neural plate to form the neural tube in vertebrates. In this case, the notochord and the floor plate are a potent source of sonic hedgehog (shh) (Jessell, 2000), and shh knockout mice exhibit a failure in neural tube closure. In these mice, the neural plate lacks characteristic bottle cells, pointing to the possibility that neural tube closure is driven by MyoII following a similar pathway to the one defined in the present study. EXPERIMENTAL PROCEDURES Genotypes Used in This Study The functional Ubi-Sqh:GFP was expressed in a sqhAX3 null mutant background (Royou et al., 2002). w, hsflp122; Tub>w[+]>Gal4/UASmYFP:myosinDN (Dawes-Hoang et al., 2005). y,w,FRT101 sqh1/UbiGFP FRT101; eyflp/+ (Karess et al., 1991). sqh1 is a hypomorphic mutation in the MRLC. rok2 FRT19A/FRT19A arm-LacZ; eyflp/+ (Winter et al., 2001). arm8 FRT19A/FRT19A arm-LacZ; eyflp/+ (Uemura et al., 1996). w, eyflp; madB1 FRT40A/FRT40A arm-LacZ and w, eyflp; mad12 FRT40A/FRT40A arm-LacZ (Wiersdorff et al., 1996). mad12 is

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an insertion in the mad locus that prevents most Dpp signaling. w, eyflp; smoD16 FRT40A/FRT40A arm-LacZ (van den Heuvel and Ingham, 1996). smoD16 is a null allele (Chen and Struhl, 1996). w, eyflp; smo3, mad1-2 FRT40A/FRT40A arm-LacZ (Curtiss and Mlodzik, 2000). mad1-2 is a hypomorphic allele, and smo3 is an amorphic allele. y, w, eyflp; [Ci+] FRT40A/FRT40A arm-LacZ;;Ci94/Ci94 (Zhang and Kalderon, 2001). y, w, eyflp; [Ci+], mad1-2 FRT40A/FRT40A armLacZ;;Ci94/Ci94 (Fu and Baker, 2003). w, eyflp; dia5 FRT40A/FRT40A arm-LacZ (Afshar et al., 2000). w, eyflp; chic221 FRT40A/FRT40A arm-LacZ (Verheyen and Cooley, 1994). The chic221 allele is an amorphic allele. w, eyflp; RhoA720M1 FRT40A/FRT40A arm-LacZ, Minutes (Strutt et al., 1997). w, eyflp; scarD37 FRT40A/FRT40A arm-LacZ (Zallen et al., 2002). w, hsflp122; ptcS2 FRT42D/FRT42D arm-LacZ (Jiang and Struhl, 1995). ptcS2 is an amorphic allele. w, hsflp122; tsr1 FRT42D/ FRT42D arm-LacZ (Gunsalus et al., 1995). w, hsflp122; ena23 FRT42D/FRT42D arm-LacZ (Ahern-Djamali et al., 1998). w, eyflp; FRT80A mbsT541/FRT80A arm-LacZ and w, eyflp; FRT80 mbsT791/ FRT80 arm-LacZ (Lee and Treisman, 2004). w, eyflp; FRT80A cul116E/FRT80A arm-LacZ and w, eyflp; FRT80A cul1107D/FRT80A arm-LacZ (Ou et al., 2002). w, eyflp; FRT80A Abl4/FRT80A arm-LacZ (Bennett and Hoffmann, 1992). w, eyflp; FRT82B ssh26-1/FRT82B arm-LacZ as well as w, eyflp; FRT82B ssh1-11/FRT82B arm-LacZ and w, eyflp; FRT82B sshP01207/FRT82B arm-LacZ (Niwa et al., 2002). w, eyflp; FRT82B wsp1/FRT82B arm-LacZ and w, eyflp; FRT82B wsp3/ FRT82B arm-LacZ (Ben-Yaacov et al., 2001). w, eyflp; FRT82B pak16/FRT82B arm-LacZ (Hing et al., 1999). pak16 is an amorphic allele. Homozygous RhoGEF24.1/RhoGEF2PX6 animals (Barrett et al., 1997). RhoGEF24.1 is a null allele, and RhoGEF2PX6 is a hypomorph. w, UAS-GFP, eyflp; TubGal4; FRT82B, TubGal80/FRT82B UAS-PakAID (Conder et al., 2004). w, UAS-GFP, eyflp; TubGal4; FRT82B, TubGal80/FRT82B, UAS-ROKCAT (Winter et al., 2001). y,w, hflp122; Tub>GFP,y+>Gal4; UAS-sqh-E20E21. UAS-sqh-E20E21 contains the sqh transgene with two activating phosphomimetic mutations (Winter et al., 2001) under the control of the UAS sequence. y,w, hflp122; Tub>GFP,y+>Gal4; UAS-spastin:GFP (Sherwood et al., 2004). Fly cultures and crosses were carried out at 25 C. Overexpression of SqhE20E21 was performed at 29 C. Immunostaining Immunostainings were performed with the following antibodies: antiArmadillo (Riggleman et al., 1990) (mouse, 1/50); anti-MyoII (Kiehart and Feghali, 1986) (rabbit, 1/500); anti-PSer19-MRLC (rabbit, 1/10; Cell Signaling Technology, Beverly, MA); anti-Diaphanous (Afshar et al., 2000) (rabbit, 1/200); anti-P-Mad (Gift from G. Morata) (rabbit, 1/500); anti-b-galactosidase (rabbit, 1/5000; Cappel Laboratories, Durham, NC); anti-b-galactosidase (goat, 1/500; Cappel Laboratories); anti-a-tubulin (Sigma, mouse, 1/2000); anti-Discs large (mouse, 1/50); phalloidin-Texas red (Sigma, 20 mM); anti-mouse, -rabbit, and -goat secondary antibodies (Jackson) (used at 1/200). Spastin:GFP-expressing cells were generated by heat shocking third-instar larvae for 30 minutes at 37 C, and the corresponding animals were dissected 5 hr later. ptc-mutant clones were induced in early second-instar larvae by performing a 1 hr heat shock at 37 C. The corresponding animals were dissected 24, 48, 72, or 96 hr after heat shock. Imaginal discs were dissected in 13 PBS and fixed in 4% formaldehyde for 20 min at room temperature. Samples were permeabilized in PBS and 0.3% Triton and then blocked in 5% FCS. Antibodies were incubated in PBS, Triton (0.3%). Imaging was performed by using a Leica SP2, SP5 or Biorad Radiance 1040 confocal microscope. Quantification Images were analyzed by using IMAGEJ v1.33. To avoid complications with variations from preparation to preparation, we chose to compare wild-type and mutant tissue from the same disc. Surface areas of wildtype and mutant patches of cells in the MF (or anterior to the MF in the case of ptc-mutant clones) were measured. For F-actin quantity, aver-

age pixel intensity over a transect in the MF (or in clones in the anterior compartment for ptc) was calculated. The data were compared by using a 2-tailed Student’s paired-t test to compare the mutant and wild-type cell responses. Supplemental Data Supplemental Data include statistical analyses and Figure S1 and are available at http://www.developmentalcell.com/cgi/content/full/13/5/ 730/DC1/. ACKNOWLEDGMENTS This work was funded by the Medical Research Council (MRC). We particularly acknowledge L. Alphey for the kind gift of the UASsqhE20E21 flies and Matthew Freeman for sharing unpublished data concerning this transgene. The authors would also like to acknowledge Buzz Baum, Nick Baker, Kathy Barrett, Andrea Brand, Roger Karess, Gines Morata, and Jessica Treisman for kindly providing us with fly stocks and reagents; the Bloomington Stock Center for fly stocks; and the Developmental Study Hybridoma Bank (DSHB) for antibodies. We are grateful to the Pichaud lab for their input during this work. R.F.W. is a recipient of an MRC career development fellowship. P.F. is a recipient of a European Molecular Biology Organization long-term fellowship. Received: February 23, 2007 Revised: July 17, 2007 Accepted: September 25, 2007 Published: November 5, 2007 REFERENCES Afshar, K., Stuart, B., and Wasserman, S.A. (2000). Functional analysis of the Drosophila diaphanous FH protein in early embryonic development. Development 127, 1887–1897. Ahern-Djamali, S.M., Comer, A.R., Bachmann, C., Kastenmeier, A.S., Reddy, S.K., Beckerle, M.C., Walter, U., and Hoffmann, F.M. (1998). Mutations in Drosophila enabled and rescue by human vasodilatorstimulated phosphoprotein (VASP) indicate important functional roles for Ena/VASP homology domain 1 (EVH1) and EVH2 domains. Mol. Biol. Cell 9, 2157–2171. Amano, M., Ito, M., Kimura, K., Fukata, Y., Chihara, K., Nakano, T., Matsuura, Y., and Kaibuchi, K. (1996). Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). J. Biol. Chem. 271, 20246–20249. Barrett, K., Leptin, M., and Settleman, J. (1997). The Rho GTPase and a putative RhoGEF mediate a signalling pathway for the cell shape changes in Drosophila gastrulation. Cell 91, 905–915. Ben-Yaacov, S., Le Borgne, R., Abramson, I., Schweisguth, F., and Schejter, E.D. (2001). Wasp, the Drosophila Wiskott-Aldrich syndrome gene homologue, is required for cell fate decisions mediated by Notch signalling. J. Cell Biol. 152, 1–13. Bennett, R.L., and Hoffmann, F.M. (1992). Increased levels of the Drosophila Abelson tyrosine kinase in nerves and muscles: subcellular localization and mutant phenotypes imply a role in cell-cell interactions. Development 116, 953–966. Borod, E.R., and Heberlein, U. (1998). Mutual regulation of decapentaplegic and hedgehog during the initiation of differentiation in the Drosophila retina. Dev. Biol. 197, 187–197. Chang, F., Drubin, D., and Nurse, P. (1997). cdc12p, a protein required for cytokinesis in fission yeast, is a component of the cell division ring and interacts with profilin. J. Cell Biol. 137, 169–182. Chanut, F., and Heberlein, U. (1997). Role of decapentaplegic in initiation and progression of the morphogenetic furrow in the developing Drosophila retina. Development 124, 559–567.

740 Developmental Cell 13, 730–742, November 2007 ª2007 Elsevier Inc.

Developmental Cell Hh Signaling and Cell Constriction

Chen, Y., and Struhl, G. (1996). Dual roles for patched in sequestering and transducing Hedgehog. Cell 87, 553–563. Conder, R., Yu, H., Ricos, M., Hing, H., Chia, W., Lim, L., and Harden, N. (2004). dPak is required for integrity of the leading edge cytoskeleton during Drosophila dorsal closure but does not signal through the JNK cascade. Dev. Biol. 276, 378–390. Costa, M., Wilson, E.T., and Wieschaus, E. (1994). A putative cell signal encoded by the folded gastrulation gene coordinates cell shape changes during Drosophila gastrulation. Cell 76, 1075–1089. Curtiss, J., and Mlodzik, M. (2000). Morphogenetic furrow initiation and progression during eye development in Drosophila: the roles of decapentaplegic, hedgehog and eyes absent. Development 127, 1325– 1336. Davidson, L.A., Koehl, M.A., Keller, R., and Oster, G.F. (1995). How do sea urchins invaginate? Using biomechanics to distinguish between mechanisms of primary invagination. Development 121, 2005–2018. Dawes-Hoang, R.E., Parmar, K.M., Christiansen, A.E., Phelps, C.B., Brand, A.H., and Wieschaus, E.F. (2005). folded gastrulation, cell shape change and the control of myosin localization. Development 132, 4165–4178. Dean, S.O., Rogers, S.L., Stuurman, N., Vale, R.D., and Spudich, J.A. (2005). Distinct pathways control recruitment and maintenance of myosin II at the cleavage furrow during cytokinesis. Proc. Natl. Acad. Sci. USA 102, 13473–13478. Edwards, K.A., and Kiehart, D.P. (1996). Drosophila nonmuscle myosin II has multiple essential roles in imaginal disc and egg chamber morphogenesis. Development 122, 1499–1511. Foe, V.E., Field, C.M., and Odell, G.M. (2000). Microtubules and mitotic cycle phase modulate spatiotemporal distributions of F-actin and myosin II in Drosophila syncytial blastoderm embryos. Development 127, 1767–1787. Fox, D.T., and Peifer, M. (2007). Abelson kinase (Abl) and RhoGEF2 regulate actin organization during cell constriction in Drosophila. Development 134, 567–578. Fu, W., and Baker, N.E. (2003). Deciphering synergistic and redundant roles of Hedgehog, Decapentaplegic and Delta that drive the wave of differentiation in Drosophila eye development. Development 130, 5229–5239. Giansanti, M.G., Bonaccorsi, S., Williams, B., Williams, E.V., Santolamazza, C., Goldberg, M.L., and Gatti, M. (1998). Cooperative interactions between the central spindle and the contractile ring during Drosophila cytokinesis. Genes Dev. 12, 396–410. Gibson, M.C., and Perrimon, N. (2005). Extrusion and death of DPP/ BMP-compromised epithelial cells in the developing Drosophila wing. Science 307, 1785–1789. Glotzer, M. (2001). Animal cell cytokinesis. Annu. Rev. Cell Dev. Biol. 17, 351–386. Gunsalus, K.C., Bonaccorsi, S., Williams, E., Verni, F., Gatti, M., and Goldberg, M.L. (1995). Mutations in twinstar, a Drosophila gene encoding a cofilin/ADF homologue, result in defects in centrosome migration and cytokinesis. J. Cell Biol. 131, 1243–1259. Hacker, U., and Perrimon, N. (1998). DRhoGEF2 encodes a member of the Dbl family of oncogenes and controls cell shape changes during gastrulation in Drosophila. Genes Dev. 12, 274–284. Hickson, G.R., Echard, A., and O’Farrell, P.H. (2006). Rho-kinase controls cell shape changes during cytokinesis. Curr. Biol. 16, 359–370. Hing, H., Xiao, J., Harden, N., Lim, L., and Zipursky, S.L. (1999). Pak functions downstream of Dock to regulate photoreceptor axon guidance in Drosophila. Cell 97, 853–863. Hooper, J.E., and Scott, M.P. (1989). The Drosophila patched gene encodes a putative membrane protein required for segmental patterning. Cell 59, 751–765.

Ishizaki, T., Morishima, Y., Okamoto, M., Furuyashiki, T., Kato, T., and Narumiya, S. (2001). Coordination of microtubules and the actin cytoskeleton by the Rho effector mDia1. Nat. Cell Biol. 3, 8–14. Jessell, T.M. (2000). Neuronal specification in the spinal cord: inductive signals and transcriptional codes. Nat. Rev. Genet. 1, 20–29. Jiang, J., and Struhl, G. (1995). Protein kinase A and hedgehog signalling in Drosophila limb development. Cell 80, 563–572. Karess, R.E., Chang, X.J., Edwards, K.A., Kulkarni, S., Aguilera, I., and Kiehart, D.P. (1991). The regulatory light chain of nonmuscle myosin is encoded by spaghetti-squash, a gene required for cytokinesis in Drosophila. Cell 65, 1177–1189. Kiehart, D.P., and Feghali, R. (1986). Cytoplasmic myosin from Drosophila melanogaster. J. Cell Biol. 103, 1517–1525. Kiehart, D.P., Ketchum, A., Young, P., Lutz, D., Alfenito, M.R., Chang, X.J., Awobuluyi, M., Pesacreta, T.C., Inoue, S., Stewart, C.T., et al. (1990). Contractile proteins in Drosophila development. Ann. N Y Acad. Sci. 582, 233–251. Kimberly, E.L., and Hardin, J. (1998). Bottle cells are required for the initiation of primary invagination in the sea urchin embryo. Dev. Biol. 204, 235–250. Kolsch, V., Seher, T., Fernandez-Ballester, G.J., Serrano, L., and Leptin, M. (2007). Control of Drosophila gastrulation by apical localization of adherens junctions and RhoGEF2. Science 315, 384–386. Lee, A., and Treisman, J.E. (2004). Excessive Myosin activity in mbs mutants causes photoreceptor movement out of the Drosophila eye disc epithelium. Mol. Biol. Cell 15, 3285–3295. Lum, L., and Beachy, P.A. (2004). The Hedgehog response network: sensors, switches, and routers. Science 304, 1755–1759. Machesky, L.M., Atkinson, S.J., Ampe, C., Vandekerckhove, J., and Pollard, T.D. (1994). Purification of a cortical complex containing two unconventional actins from Acanthamoeba by affinity chromatography on profilin-agarose. J. Cell Biol. 127, 107–115. Methot, N., and Basler, K. (1999). Hedgehog controls limb development by regulating the activities of distinct transcriptional activator and repressor forms of Cubitus interruptus. Cell 96, 819–831. Miki, H., and Takenawa, T. (2003). Regulation of actin dynamics by WASP family proteins. J. Biochem. (Tokyo) 134, 309–313. Morize, P., Christiansen, A.E., Costa, M., Parks, S., and Wieschaus, E. (1998). Hyperactivation of the folded gastrulation pathway induces specific cell shape changes. Development 125, 589–597. Nakagawa, H., Miki, H., Ito, M., Ohashi, K., Takenawa, T., and Miyamoto, S. (2001). N-WASP, WAVE and Mena play different roles in the organization of actin cytoskeleton in lamellipodia. J. Cell Sci. 114, 1555–1565. Nikolaidou, K.K., and Barrett, K. (2004). A Rho GTPase signalling pathway is used reiteratively in epithelial folding and potentially selects the outcome of Rho activation. Curr. Biol. 14, 1822–1826. Niwa, R., Nagata-Ohashi, K., Takeichi, M., Mizuno, K., and Uemura, T. (2002). Control of actin reorganization by Slingshot, a family of phosphatases that dephosphorylate ADF/cofilin. Cell 108, 233–246. Ou, C.Y., Lin, Y.F., Chen, Y.J., and Chien, C.T. (2002). Distinct protein degradation mechanisms mediated by Cul1 and Cul3 controlling Ci stability in Drosophila eye development. Genes Dev. 16, 2403–2414. Palazzo, A.F., Cook, T.A., Alberts, A.S., and Gundersen, G.G. (2001). mDia mediates Rho-regulated formation and orientation of stable microtubules. Nat. Cell Biol. 3, 723–729. Parks, S., and Wieschaus, E. (1991). The Drosophila gastrulation gene concertina encodes a G a-like protein. Cell 64, 447–458. Ready, D.F., Hanson, T.E., and Benzer, S. (1976). Development of the Drosophila retina, a neurocrystalline lattice. Dev. Biol. 53, 217–240. Riggleman, B., Schedl, P., and Wieschaus, E. (1990). Spatial expression of the Drosophila segment polarity gene armadillo is posttranscriptionally regulated by wingless. Cell 63, 549–560.

Developmental Cell 13, 730–742, November 2007 ª2007 Elsevier Inc. 741

Developmental Cell Hh Signaling and Cell Constriction

Royou, A., Sullivan, W., and Karess, R. (2002). Cortical recruitment of nonmuscle myosin II in early syncytial Drosophila embryos: its role in nuclear axial expansion and its regulation by Cdc2 activity. J. Cell Biol. 158, 127–137. Schroeder, T.E. (1970). Neurulation in Xenopus laevis. An analysis and model based upon light and electron microscopy. J. Embryol. Exp. Morphol. 23, 427–462. Seher, T.C., Narasimha, M., Vogelsang, E., and Leptin, M. (2006). Analysis and reconstitution of the genetic cascade controlling early mesoderm morphogenesis in the Drosophila embryo. Mech. Dev. 124, 167– 179.

Verheyen, E.M., and Cooley, L. (1994). Profilin mutations disrupt multiple actin-dependent processes during Drosophila development. Development 120, 717–728. Wasserman, S. (1998). FH proteins as cytoskeletal organizers. Trends Cell Biol. 8, 111–115. Watanabe, N., Madaule, P., Reid, T., Ishizaki, T., Watanabe, G., Kakizuka, A., Saito, Y., Nakao, K., Jockusch, B.M., and Narumiya, S. (1997). p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J. 16, 3044–3056.

Shen, J., and Dahmann, C. (2005). Extrusion of cells with inappropriate Dpp signalling from Drosophila wing disc epithelia. Science 307, 1789– 1790.

Watanabe, N., Kato, T., Fujita, A., Ishizaki, T., and Narumiya, S. (1999). Cooperation between mDia1 and ROCK in Rho-induced actin reorganization. Nat. Cell Biol. 1, 136–143.

Sherwood, N.T., Sun, Q., Xue, M., Zhang, B., and Zinn, K. (2004). Drosophila spastin regulates synaptic microtubule networks and is required for normal motor function. PLoS Biol. 2, e429.

Wiersdorff, V., Lecuit, T., Cohen, S.M., and Mlodzik, M. (1996). Mad acts downstream of Dpp receptors, revealing a differential requirement for dpp signalling in initiation and propagation of morphogenesis in the Drosophila eye. Development 122, 2153–2162.

Somogyi, K., and Rorth, P. (2004). Evidence for tension-based regulation of Drosophila MAL and SRF during invasive cell migration. Dev. Cell 7, 85–93. Strutt, D.I., Weber, U., and Mlodzik, M. (1997). The role of RhoA in tissue polarity and Frizzled signalling. Nature 387, 292–295. Tsuji, T., Ishizaki, T., Okamoto, M., Higashida, C., Kimura, K., Furuyashiki, T., Arakawa, Y., Birge, R.B., Nakamoto, T., Hirai, H., and Narumiya, S. (2002). ROCK and mDia1 antagonize in Rho-dependent Rac activation in Swiss 3T3 fibroblasts. J. Cell Biol. 157, 819–830. Uemura, T., Oda, H., Kraut, R., Hayashi, S., Kotaoka, Y., and Takeichi, M. (1996). Zygotic Drosophila E-cadherin expression is required for processes of dynamic epithelial cell rearrangement in the Drosophila embryo. Genes Dev. 10, 659–671. van den Heuvel, M., and Ingham, P.W. (1996). smoothened encodes a receptor-like serpentine protein required for hedgehog signalling. Nature 382, 547–551.

Winter, C.G., Wang, B., Ballew, A., Royou, A., Karess, R., Axelrod, J.D., and Luo, L. (2001). Drosophila Rho-associated kinase (Drok) links Frizzled-mediated planar cell polarity signalling to the actin cytoskeleton. Cell 105, 81–91. Wolff, T., and Ready, D.F. (1991). The beginning of pattern formation in the Drosophila compound eye: the morphogenetic furrow and the second mitotic wave. Development 113, 841–850. Xu, T., and Rubin, G.M. (1993). Analysis of genetic mosaics in developing and adult Drosophila tissues. Development 117, 1223–1237. Zallen, J.A., Cohen, Y., Hudson, A.M., Cooley, L., Wieschaus, E., and Schejter, E.D. (2002). SCAR is a primary regulator of Arp2/3-dependent morphological events in Drosophila. J. Cell Biol. 156, 689–701. Zhang, Y., and Kalderon, D. (2001). Hedgehog acts as a somatic stem cell factor in the Drosophila ovary. Nature 410, 599–604.

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