Rok is ubiquitously expressed throughout development, in a pattern essentially identical to that of Rho1 (Mizuno, 1999).
Cell fate diversity can be achieved through the asymmetric segregation of cell fate determinants. In the Drosophila embryo, neuroblasts divide asymmetrically and in a stem cell fashion. The determinants Prospero and Numb localize in a basal crescent and are partitioned from neuroblasts to their daughters (GMCs). Nonmuscle myosin II regulates asymmetric cell division by an unexpected mechanism, excluding determinants from the apical cortex. Myosin II is activated by Rho kinase and restricted to the apical cortex by the tumor suppressor Lethal (2) giant larvae. During prophase and metaphase, myosin II prevents determinants from localizing apically. At anaphase and telophase, myosin II moves to the cleavage furrow and appears to 'push' rather than carry determinants into the GMC. Therefore, the movement of myosin II to the contractile ring not only initiates cytokinesis but also completes the partitioning of cell fate determinants from the neuroblast to its daughter (Barros, 2003).
Myosin II is activated by phosphorylation of its regulatory light chain by Rho kinase. Myosin was inactivated at the time of neuroblast cell division by inhibition of Rho kinase. Myosin II no longer localizes at the apical neuroblast cortex but instead spreads into the cytoplasm, and basal protein localization is disrupted. Although F-actin and Lgl remain uniformly at the cortex, cell fate determinants are now found around the entire cell cortex, demonstrating that apical cortical myosin is required to confine determinants to the basal half of the cell. Inhibition of Rho kinase also blocks cytokinesis, although the defect in basal protein localization is unlikely to be the consequence of mitotic arrest or a block in cytokinesis. First, basal protein localization is not disrupted in neuroblasts arrested in mitosis by colcemid treatment. Second, mitosis occurs without cytokinesis in pebble mutants, but the resultant polyploid neuroblasts still localize Numb and Prospero asymmetrically. Finally, the loss of asymmetry resulting from Rho kinase inhibition can be rescued by expression of a constitutively active form of the myosin II regulatory light chain (SqhE20E21). It is concluded that myosin II is required to restrict cell fate determinants to the basal cortex (Barros, 2003).
How does myosin II restrict neuroblast proteins to the basal side of the cell cortex? Myosin II and Miranda occupy primarily opposite sides at the neuroblast cortex: myosin II is concentrated at the apical cortex while Miranda localizes as a basal crescent. As myosin II shifts to the cleavage furrow, Miranda is segregated into the forming GMC. The apical F-actin compartment may be modified by myosin II to exclude binding of basal proteins like Miranda. Active myosin II requires Rho kinase activity and depends on inactivation of Lgl at the apical cortex by aPKC (Betschinger, 2003). Ectopic expression of a nonphosphorylatable form of Lgl, in which the conserved aPKC-dependent phosphorylation sites are mutated from Serines to Alanines (Lgl-3A), results in mislocalization of Miranda around the neuroblast cortex (Betschinger, 2003). The data support a spatially regulated interaction between myosin II and Lgl. Myosin is apically localized in wild-type neuroblasts, corresponding to the domain in which Lgl is inactivated by aPKC. In lgl mutants, myosin is no longer restricted apically but localizes around the entire cell cortex. Conversely, when nonphosphorylatable Lgl is expressed in neuroblasts, myosin is inhibited throughout the cell and drops off the cortex. It is proposed that myosin II is activated and can form filaments at the apical cortex, where phosphorylated Lgl is inactive and unable to bind myosin II. Myosin may then modify the actin cytoskeleton to prevent the binding of Miranda. At the basal cortex, in the absence of aPKC, Lgl is active and can bind and inhibit myosin. Myosin cannot form filaments, which are required for it to bind to the actin cortex. As a result, Miranda can bind to the basal cortex (Barros, 2003).
At anaphase, myosin II moves to the equator and appears to 'push' cell fate determinants into the daughter cell. This movement is regulated in an Lgl-independent fashion and occurs whether myosin is restricted to the apical cortex or is uniformly cortical (as in lgl mutants). Cortical myosin is essential, however, to efficiently segregate determinants into the GMC at telophase (telophase rescue). In neuroblasts expressing Lgl-3A, myosin II is cytoplasmic, and determinants are not partitioned to the daughter cell. Nonetheless, at telophase, myosin seems to be recruited from the cytoplasm, since it still accumulates to the cleavage furrow. Thus three separate steps of myosin regulation in neuroblasts can be defined. First, myosin forms an apical crescent. This is positively regulated by Rho kinase and negatively regulated by Lgl. Second, cortical myosin moves to the equator. This movement occurs independently of Lgl. Third, cortical and cytoplasmic myosin accumulates at the cleavage furrow, a step that is also Lgl independent. Rho Kinase activation seems to be important for all three steps of myosin II regulation. When Rho kinase is inhibited, myosin falls into the cytoplasm, and there is no cleavage furrow formation (Barros, 2003).
In conclusion, these results demonstrate that myosin II acts downstream of Lgl and the apical protein complex to regulate the segregation of cell fate determinants. Myosin II does not negatively regulate basal protein targeting, as has previously been suggested nor does it transport determinants directly. Instead, it is proposed that myosin II acts in a novel fashion, excluding determinants from the apical cortex and 'pushing' them into the GMC at anaphase and telophase. Myosin II might modify the actin cytoskeleton to prevent determinants binding, although the actual structure formed and the physical change in the actin cytoskeleton remains to be determined (Barros, 2003).
The Rho-kinases are widely utilized downstream targets of the activated Rho GTPase that have been directly implicated in many aspects of Rho-dependent effects on F-actin assembly, acto-myosin contractility, and microtubule stability, and consequently play an essential role in regulating cell shape, migration, polarity, and division. The single closely related Drosophila Rho-kinase ortholog, DRok, is required for several aspects of oogenesis, including maintaining the integrity of the oocyte cortex, actin-mediated tethering of nurse cell nuclei, 'dumping' of nurse cell contents into the oocyte, establishment of oocyte polarity, and the trafficking of oocyte yolk granules. These defects are associated with abnormalities in DRok-dependent actin dynamics and appear to be mediated by multiple downstream effectors of activated DRok that have previously been implicated in oogenesis. DRok regulates at least one of these targets, the membrane cytoskeletal cross-linker DMoesin, via a direct phosphorylation that is required to promote localization of DMoesin to the oocyte cortex. The collective oogenesis defects associated with DRok deficiency reveal its essential role in multiple aspects of proper oocyte formation and suggest that DRok defines a novel class of oogenesis determinants that function as key regulators of several distinct actin-dependent processes required for proper tissue morphogenesis (Verdier, 2006a).
DRok has been previously implicated as an effector of the DRho1 GTPase in the regulation of planar cell polarity in the eye and in the wing, downstream of Frizzled-Dishevelled signals. In germline cells, DRok and DRho1 mutants exhibit some overlapping actin defects; e.g., the oocyte cortex exhibits a more diffuse F-actin distribution in both Drok2 GLCs and Rho1 loss-of-function egg chambers in which Rho1 levels have been reduced in a heterozygous mutant Rho1 and wimp background. In addition, wild-type oocytes injected with the Rho-inhibitory C3 toxin exhibit the same ooplasmic streaming defects as Drok2 mutant oocytes, strongly suggesting that the germline clone phenotypes reflect the disruption of DRho1-DRok signaling in germ cells (Verdier, 2006a).
Previous analysis of the polarity of Dmoe GLC oocytes indicated that DMoesin is specifically required for the localization of posterior determinants such as oskar mRNA and Oskar protein but not the formation of the dorso-ventral axis nor the anterior pole. DMoesin appears to function in maintaining posterior polarity by anchoring actin to the membrane cortex which in turn anchors microtubule-delivered oskar mRNA and its protein product Oskar to the posterior pole. Similarly, in Drok2 GLCs, the localization of anterior (bicoid) or dorso-ventral determinants (gurken) is not altered although most oskar mRNA is found mislocalized within the ooplasm starting at stage 9. While the establishment of oocyte polarity generally depends upon microtubule cytoskeleton organization, it has been reported that Dmoe null mutations do not disrupt the microtubule cytoskeleton and do not perturb its polarity. Similarly, this study observed that microtubules in Drok2 mutant oocytes appear normal. Taken together with the fact that some oskar mRNA remains anchored at the posterior tip of the oocyte plasma membrane in Drok2 GLCs, as is seen in Dmoe GLCs, this indicates that oskar mislocalization, and consequently, the alteration of posterior polarity in Drok2 GLCs is not due to an abnormally organized microtubule cytoskeleton. Moreover, unlike other germline clone mutants with oocyte polarity defects, such as chic or capu, the Drok2 and Dmoe polarity defects most likely reflect the incapacity of the disorganized subcortical actin cytoskeleton to properly anchor oskar at the posterior membrane of the oocyte (Verdier, 2006a).
The similarity between the oskar polarity phenotype of Drok2 and Dmoe GLCs is also consistent with a likely role for DMoesin as an essential DRok substrate that mediates its effects on the formation of posterior polarity and further supports the functional significance of a signaling pathway from DRok to DMoesin to the actin cytoskeleton in oocyte development. Moreover, the proper localization of Gurken, as defined by the position of the oocyte nucleus, which migrates in a microtubule-dependent manner from the posterior to the anterior and then to the antero-dorsal side of the oocyte starting at stage 8, is consistent with the presence of a grossly normal appearing microtubule cytoskeleton in DRok-deficient oocytes (Verdier, 2006a).
A majority of mutations resulting in egg chambers with dorso-ventral axis patterning defects, such as dorsal appendages aberrations, have been associated with genes encoding components of the Gurken-EGFR signaling pathway. Cross-signaling between the oocyte and the surrounding follicle cells at the antero-dorsal side of the oocyte has been extensively studied and involves the binding of the secreted Gurken ligand to the EGFR present on the apical membrane of follicle cells and subsequent activation of downstream signaling to control the formation of follicle cell-derived dorsal structures. Although Gurken localization is correct in Drok2 mutant oocytes, loss of Gurken secretion (in 80% of the Drok2 mutant oocytes) in the intercellular space between the oocyte membrane and the follicle apical membranes indicates the likelihood of altered communication between the oocyte and surrounding follicle cells, possibly resulting in a disruption of the EGFR signaling pathway leading to dorsal appendage defects. The observed requirement for DRok in Gurken secretion may reflect a well established role of Rho signaling in the control of vesicular trafficking and secretion. However, it remains possible that the apparent absence of Gurken secretion into the intercellular space reflects a consequence of the observed disruption of oocyte plasma membrane integrity (Verdier, 2006a).
The unexpected observation in time-lapse confocal microscopy studies that most autofluorescent yolk granules in Drok2 mutant oocytes or Rho-inhibitory C3 transferase-treated wild-type egg chambers accumulate at the oocyte membrane suggests a role for DRho1 and DRok in early vitellogenesis. Vitellogenesis is a process that begins around stage 8 and is defined by the co-secretion of vitelline membrane and yolk material by the surrounding follicle cells leading to the eventual formation of chorionic structures of the egg and normal oocyte growth, respectively. After their secretion, yolk proteins are internalized into the oocyte through endocytosis and are swirled around the ooplasm at later stages, when microtubule-dependent streaming occurs. The high concentration of yolk granules at the oocyte membrane from early vitellogenesis underlies a possible defect in endocytosis of the yolk granules. Together with the fact that C3-treated egg chambers and Drok2 GLCs exhibit an identical yolk granule phenotype, this suggests that DRok mediates Rho1's role in the trafficking of yolk granules at the oocyte plasma membrane. In addition, nurse cells also normally accumulate yolk material and transfer it to the oocyte. The detection of yolk granules moving to the plasma membrane of Drok2 mutant oocytes or oocytes in C3-treated egg chambers after they are deposited by the nurse cells is an intriguing phenotype that has not been previously reported and may reflect a trafficking defect in the ooplasm. Further studies to examine molecular components of the endocytic machinery will be required to develop a better understanding of the roles of Rho1 and DRok in yolk granule trafficking within the ooplasm. Notably, it is also conceivable that alteration of oocyte plasma membrane integrity through disruption of actin cytoskeleton organization in most Drok2 GLCs, as was observed, could exert a secondary effect on the endocytosis of yolk granules (Verdier, 2006a).
Because of the yolk granule phenotype in Drok2 GLCs in early oogenesis, it is not possible to visualize microtubule cytoskeleton dynamics at later stages in time-lapse confocal microscopy. Thus, it is difficult to determine whether Drok2 mutant oocytes would undergo normal or premature ooplasmic streaming at stages 10b–11. As a functional relationship between actin and microtubule cytoskeletons has been suggested based on findings with several mutants with oogenesis defects, it is quite conceivable that the abnormalities of the actin cytoskeleton in Drok2 mutant oocytes could affect microtubule cytoskeleton dynamics. Indeed, it has been demonstrated that some aspect of the actin cytoskeleton normally represses microtubule-based streaming within the oocyte. Thus, it is possible that the accumulation of yolk granules near the plasma membrane of Drok2 mutant oocytes reflects a combination of trafficking/endocytosis defects and actin cytoskeleton perturbation-induced alteration of microtubule cytoskeleton dynamics in the ooplasm during early oogenesis (Verdier, 2006a).
The oocyte volume in Drok2 GLCs is frequently smaller than that seen in wild-type oocytes, before the rapid phase of cytoplasmic transport takes place. This suggests a possible defect in the slow phase of cytoplasmic transport. It has been previously reported that transport of some particles towards the oocyte during stages 7–10A depends upon a proper acto-myosin network. In addition, sqhAX3 GLCs exhibit a similar oocyte size defect. sqhAX3 is a loss-of-function mutation in the sqh locus which codes for the Drosophila ortholog of myosin light chain of myosin II. Taken together with the fact that DRok has been shown to phosphorylate Sqh in vivo, these data suggest that DRok mediates, via regulation of Sqh, some aspects of the acto-myosin contractility involved in cytoplasmic transport from early stages of oogenesis (Verdier, 2006a).
The observation of dumpless-like oversized nurse cells in most of Drok2 GLCs also supports a role for DRok in the rapid phase of cytoplasmic transport at stages 10B–11 of oogenesis. Unlike other classes of dumpless mutants including chickadee, singed or quail, failure of rapid cytoplasmic transport from the Drok2 mutant nurse cells to the oocyte does not result from the obstruction of the ring canals by unanchored nurse cell nuclei, suggesting that Drok constitutes a distinct class of dumpless-like mutants. In addition, in sqhAX3 GLCs, dumpless nurse cells are associated with a lack of acto-myosin contractility by nurse cells, as revealed by mislocalization of myosin II and by absence of the perinuclear organization of actin filaments bundles in the nurse cells. Therefore, sqhAX3 mutant nurse cells cannot contract properly to expulse their cytoplasm through otherwise weakly damaged ring canals. Drok2 and sqhAX3 mutant nurse cells do not share the same actin filament phenotype; Drok2 mutant nurse cells exhibit a more dramatic phenotype associated with absence of radial filaments and disorganization of cortical actin. It is, however, likely that DRok and Sqh are part of the same signaling pathway that regulates acto-myosin contractility in nurse cells; it has already been shown that DRok phosphorylates Sqh in Drosophila development. Moreover, the severity of the Drok2 mutant F-actin phenotypes may reflect DRok's potential to engage multiple distinct downstream substrates, of which Sqh is only one. Significantly, the actin-binding protein, adducin, is also reportedly a direct substrate for mammalian Rho-kinases, and the Drosophila Adducin ortholog, Hts, is a major component of ring canals. Thus, it is possible that the observed defects in ring canal morphology in Drok2 GLCs involve abnormal regulation of adducin by DRok. However, it is difficult to determine whether this ring canal phenotype contributes to the dumpless-like nurse cell phenotype observed in Drok2 GLCs (Verdier, 2006a).
The observation that nurse cell nuclei are substantially increased in size in Drok2 GLCs suggests a possible involvement of DRok in increased endoreplication of the nurse cells. The Rho-related Rac and Cdc42 GTPases have previously been associated with endoreplication in porcine aortic endothelial (PAE) cells, although Rho has not been implicated thus far. Interestingly, this nurse cell nuclei phenotype has not been observed in other previously described GLC mutants of other actin cytoskeleton-regulating signaling components that exhibit oogenesis defects. Thus, chic as well as sqhAX3 GLCs reveal cytokinesis defects associated with the presence of multinucleated nurse cells. In addition, the majority of sqhAX3 mutant egg chambers harbor less than 15 nurse cells (64% of sqhAX3 mutant egg chambers have less than 7 nurse cells), a phenotype that is not shared by Drok2 mutant nurse cells. These findings also suggest that Drok2 defines a new category of oogenesis mutants that affect the actin cytoskeleton (Verdier, 2006a).
Both Dmoe and Drok2 GLCs exhibit similar actin defects in the oocyte, associated with a loose uneven cortical actin distribution and the presence of actin clumps in the ooplasm and near the cortex. Moreover, phospho-DMoesin levels are decreased at the cortex or mislocalized within the ooplasm of Drok2 GLCs and the conserved kinase domain of Rho-kinase phosphorylates DMoesin on threonine 559 in vitro. A potential mechanism for the DRok-DMoesin signal in this setting is that DRok controls actin reorganization through phosphorylation of DMoesin, which has been previously shown to cross-link actin to the plasma membrane when phosphorylated on T559 at the oocyte cortex. However, the detection of some phospho-DMoesin in the Drok2 GLCs indicates that the critical T559 residue can be phosphorylated by other kinases in the oocyte. Indeed, direct phosphorylation of T559 of mammalian Moesin by protein kinase C (PKC)-θ has been shown in vitro. In addition, mammalian Rho-kinase and PAK have been reported to both phosphorylate the very conserved T508 residue of LIM-kinase in vitro. Therefore, phosphorylation of the conserved T559 residue of Moesin by additional kinases might also occur in Drosophila, highlighting the complexity of cross-talk within developmental signaling pathways (Verdier, 2006a).
The observation that Drok2 mutant oocytes are morphologically more affected than Dmoe mutant oocytes with regard to the deformed plasma membrane also suggests that to exert its functions at the oocyte cortex, DRok is not only signaling to DMoesin but probably also to additional downstream targets that cooperate with DMoesin in the maintenance of the cortical actin cytoskeleton. The strong phenotype associated with the deformed oocyte plasma membrane, which separates dramatically from the apical plasma membranes of the follicle cell layer in most Drok2 GLCs, raises an intriguing question about DRok's apparent role in an adhesive process. That specific phenotype has not been previously reported in studies of other oogenesis mutants associated with defective adhesion between the oocyte and the surrounding follicle cells. Previous reports regarding such adhesion largely address cross-signaling between the apical Notch receptor and the germline-derived putative secreted and transmembrane proteins, Brainiac and Egghead, respectively, in which germline loss of either Brainiac or Egghead results in loss of epithelial apico-basal polarity and accumulation of follicular epithelial cells in multiple layers around the oocyte, but does not lead to a physical separation between the oocyte and the follicle cells membranes. The unique phenotype of Drok2 GLCs could reflect a role for DRok in mediating a distinct signaling pathway from the oocyte to regulate its shape and its adherence to the surrounding follicle cells. Alternatively, the aberrant morphology of the nurse cells, which appear to 'push' against the oocyte without contracting, might produce a mechanical stress on the oocyte itself that prevents it from remaining apposed to the follicle cell layer. Notably, it was found that the follicle cells themselves also appear to require DRok function for the maintenance of their shape, and it is possible that their ability to signal to the oocyte is also affected by DRok deficiency (Verdier, 2006a).
In summary, the single closely related Drosophila Rho-kinase ortholog, DRok, is required for several aspects of oogenesis, including maintaining the integrity of the oocyte cortex, actin-dependent tethering of nurse cell nuclei, 'dumping' of nurse cell contents into the oocyte, establishment of oocyte polarity, and the trafficking of oocyte yolk granules. It is likely that several previously identified direct phosphorylation targets of DRok, including DMoesin, Sqh (myosin light chain), and Hts (adducin), which have each been implicated in various aspects of oogenesis, mediate at least some of the functions of DRok in developing egg chambers. These findings indicate an essential role for Rho-DRok signaling via multiple DRok effectors in several distinct aspects of oogenesis (Verdier, 2006a).
The Rho-kinases (ROCKs) are major effector targets of the activated Rho GTPase that have been implicated in many of the Rho-mediated effects on cell shape and movement via their ability to affect acto-myosin contractility. The role of ROCKs in cell shape change and motility suggests a potentially important role for Rho-ROCK signaling in tissue morphogenesis during development. Indeed, in Drosophila, a single ROCK ortholog, DRok, has been identified and has been found to be required for establishing planar cell polarity. A potential role for DRok in additional aspects of tissue morphogenesis was examined using an activated form of the protein in transgenic flies. The findings demonstrate that DRok activity can influence multiple morphogenetic processes, including eye and wing development. Furthermore, genetic studies reveal that Drok interacts with multiple downstream effectors of the Rho GTPase signaling pathway, including non-muscle myosin heavy chain, adducin, and Diaphanous in those developmental processes. Finally, in overexpression studies, it was determined that Drok and Drosophila Lim-kinase interact in the developing nervous system. These findings indicate widespread diverse roles for DRok in tissue morphogenesis during Drosophila development, in which multiple DRok substrates appear to be required (Verdier, 2006b; full text of article).
Among the Rho-kinase substrates that have been strongly implicated in neural development are the Lim-kinases. The single Drosophila Lim-kinase (DLimk) is required for proper synapse formation and proper regulation of its activity is necessary for normal axon growth. To determine whether DRok-mediated activation of DLimk plays a role in proper neural development, transgenic flies expressing activated DRok were crossed with flies over-expressing DLimk to examine phenotypes in the developing nervous system. First, using GMR-driven transgenes to identify a potential interaction in the developing eye, it was observed that while overexpression of DLimk causes no detectable effects on eye development, co-expression of DLimk with activated DRok results in a dramatic disruption of eye development associated with a severe morphology defect of the external eye and a reduced overall eye size. Since the effects of a single-copy DRok-cat transgene on exterior eye structures in this setting are relatively mild, this finding is consistent with a synergistic interaction between these two proteins, and suggests that a DRok-DLimk signal may be influencing normal development. Second, a similar synergistic interaction between DRok and DLimk was observed in the developing central nervous system. Using an elav-GAL4 driver to express UAS-linked Drok-cat and Dlimk in developing neurons, it was observed that neither protein alone causes any detectable effect on the appearance of the embryonic nervous system, whereas co-expression of the proteins results in the appearance of breaks along the ventral nerve cord. These findings suggest that DLimk is likely to mediate at least some of the DRok-dependent functions in the developing nervous system (Verdier, 2006b).
In conclusion, genetic analysis of DRok in development, using ovexpression studies in the eye, in the wing and in the CNS indicates that stringent regulation of DRok activity is required for various developmental processes, such as photoreceptor maintenance and wing vein formation. In addition, the overexpression system has revealed zipper, the Drosophila nonmuscle myosin heavy chain, as a strong genetic interactor of DRok, as seen in other reported developmental events such as dorsal closure and wing planar cell polarity, confirming that myosin II is a key downstream mediator of Rho-kinase biological effects in several morphogenetic processes. Moreover, DRok interacts with another target protein, DLimk, to influence some other aspects of tissue morphogenesis, including CNS development (Verdier, 2006b).
The Drosophila egg chamber is an organ composed of a somatic epithelium that covers a germline cyst. After egg-chamber formation, the germline cells grow rapidly without dividing while the surface of the epithelium expands by cell proliferation. The mechanisms that coordinate growth and morphogenesis of the two tissues are not known. This study identifies a role for the actomyosin cytoskeleton in this process. Myosin activity is restricted to the epithelium's apical surface, which is facing the growing cyst. The epithelium collapses in the absence of myosin activity; the force that deforms the epithelium originates from the growing cyst. Thus, myosin activity maintains epithelial shape by balancing the force emanating from cyst growth. Further, these data indicate that cyst growth induces cell division in the epithelium. In addition, apical restriction of myosin activity is controlled. Myosin is activated at the apical cortex by localized Rho kinase and inhibited at the basolateral cortex by PP1β9C. In addition, these data indicate that active myosin is apically anchored by the Bazooka/Par-6/aPKC complex (Wang, 2007).
To analyze the correlation between cyst growth and follicle cell division, dividing cells in the follicular epithelium were counted. Within the first 56 hr that are required to form a stage 3 egg chamber, cell-division rates are very low. In the 14 hr period between stage 4 and stage 6, however, cell-division rates continuously increase. During this time, the volume of the cyst increases approximately 11-fold. This parallel increase in mitosis and cyst growth reflects how the growth of the inner cyst is compensated by cell division in the outer follicular epithelium. After stage 7, the follicle cells stop dividing and undergo diverse morphological changes (Wang, 2007).
Newly formed egg chambers are round and change their shape to ellipsoid during early oogenesis. After stage 7, the process of egg-chamber elongation, which is mediated by a polarized actin cytoskeleton within the follicular epithelium, starts. Actin fibers at the basal cortex of the follicle cells run perpendicular to the anterior-posterior axis of the egg chamber, and their contraction leads to an axis expansion. The mechanisms that shape the egg chamber before elongation takes place are unknown. The simultaneous and rapid growth of cyst and epithelium after stage 3 indicates that the development of the two tissues is precisely coordinated. It is, however, unclear how epithelial morphogenesis and proliferation are coupled to the growth of the cyst (Wang, 2007).
The actin cytoskeleton is central for the cell shape, and is thus a possible candidate involved in a controlled epithelial response to the cyst growth. The activity in the actomyosin cytoskeleton was examined by using an antibody specific for the activated form of the regulatory light chain of nonmuscle myosin II (RMLC; Spaghetti squash). The phospho-specific RMLC antibody binds to phosphoSerine21 and reveals myosin in its active state. Around stage 3 of oogenesis, myosin activity restricts to the apical cortex of the follicle cells, where it is maintained until late oogenesis. After stage 7, myosin activity is also present at the basal cortex of the follicle cells in the actin bundles required for egg chamber elongation. Optical confocal sections reveal a pattern of myosin activity in these long parallel bundles that is reminiscent of stress fibers. In contrast, at the apical cortex, myosin is active in short fibers with random orientation reminiscent of a web (Wang, 2007).
The membrane domains of the follicle cells are established before myosin activity restricts to the apical cortex at stage 3. To examine how apical myosin activation relates to follicle cell polarity, mutants affecting epithelial polarity were examined. To avoid perdurance of the wild-type protein after clone induction, focus was placed on large clones, or clones spanning the whole epithelium. The adherence junctions are central for the organization of the apical actin cytoskeleton, and the domain of myosin activity extends into the region where they localize. Therefore null mutants of armadillo (arm), which encodes Drosophila β-catenin, were examined. It has been shown that the adherence junctions are disrupted in arm follicle cell clones since neither DE- or DN-cadherin are detectable. As a result, arm mutant cells exhibit strong cell-shape defects. Surprisingly, it was found that myosin activity is clearly restricted to the apical membrane in arm follicle cell clones. Thus, myosin activity restricts apically in the absence of adherence junctions (Wang, 2007).
The apical membrane domain is established by the Crumbs (Crb)/Stardust (Sdt)/Patj complex and the Bazooka (Baz)/Par-6/aPKC complex. All these proteins localize, like pRMLC, to the apical membrane of the follicular epithelium. In epithelia lacking crb, myosin restriction is affected as revealed by the interrupted apical pRMLC pattern and by ectopic activity at the basal membrane. However, apical myosin activity is not completely disrupted as broad regions of the epithelium still concentrate higher levels of pRMLC at the apical compared to the basal cortex. In contrast, par-6, aPKC and baz mutants abolish the formation of the apical myosin activity. In these mutants, apical pRMLC restriction is lost, and ectopic myosin activity is detectable in the cytoplasm and at the basal cortex. To test whether the two apical complexes cooperate in apical myosin restriction, baz sdt double mutants were examined. The phenotype of the double mutants is, however, very similar to that of the baz single mutants, suggesting that apical myosin activity is controlled by the Baz/Par-6/aPKC complex (Wang, 2007).
To further analyze this interaction, the Baz/Par-6/aPKC complex was immunoprecipitated from ovaries using an antibody against Baz. Western-blot analysis of the precipitated protein complex reveals a strong enrichment of Baz and aPKC. Notably, pRMLC is also present in the precipitated protein complex, indicating an association of Baz and active myosin. Taken together, these genetic data show that baz, par-6, and aPKC are required for apical myosin restriction, and biochemical data show that Baz associates with pRMLC. This suggests the Baz/Par-6/aPKC complex anchors active myosin at the apical cortex. To further analyze the role of the complex in the apical restriction of myosin activity, its localization was examined in mutants that affect pRMLC localization. Consistent with a function in the anchoring of active myosin, it was found that apical aPKC localization is not affected in arm mutants, in which pRMLC is apically restricted. Further, apical aPKC localization is interrupted in crb mutants, in which pRMLC localization is also interrupted. In summary, the data suggest that the Baz/Par-6/aPKC complex anchors active myosin at the apical cortex independently of the adherence junctions (Wang, 2007).
To examine how myosin activity is inhibited during early oogenesis at the basal and lateral cortex, the localization and function of PP1β9C, the phosphatase that deactivates phosphorylated RMLC. was examined. In follicle cells, PP1β9C is ubiquitously distributed as revealed by a hemagglutinin (HA) fusion protein. PP1β9C is encoded by flap wing (flw). Western-blot analysis of the viable flw1 allele showed that the total pRMLC levels in ovaries are increased 2.8-fold compared to those of the wild-type. The total increase is the result of ectopic myosin activity in the follicular epithelium. This is revealed by flw6 follicle cell clones and homozygous flw1 mutant egg chambers, which show pRMLC staining at the basal and lateral cortex. Interestingly, the ectopic Myosin activity is accompanied by an irregular and wavy appearance of the apical surface of the epithelium. In addition, flw mutant egg chambers are not round or ellipsoid like the wild-type but are either stretched or develop bulges. The coincidence of the altered shape with the ectopic pRMLC staining in the follicular epithelium suggests that the abnormal shape is the result of ectopic myosin activity. This is confirmed by the finding that the expression of constitutively active RMLC results in a very similar phenotype. The defects in flw mutants are not secondary effects of mislocalization of the Baz/Par-6/aPKC complex as the localization of aPKC is indistinguishable from that of the wild-type. In summary, these results show that PP1β9C activity is required to prevent myosin activity at the basal and lateral cortex. They further suggest that during early oogenesis, myosin activity has to be restricted to the apical cortex to ensure the development of normally shaped egg chambers (Wang, 2007).
To investigate how myosin is activated at the apical cortex, the function of Rok, which has been shown to regulate myosin phosphorylation, was analyzed. Myosin phosphorylation is greatly reduced but still detectable in rok mutant follicle cell clones. This confirms that Rok phosphorylates myosin in the follicular epithelium, but also indicates that Rok is not the only kinase involved in myosin activation. A HA-Rok fusion protein accumulates in particles at the apical cortex of the follicle cells, which are in close proximity to the web-like myosin fibers. Thus, localized Rok activates myosin in the follicular epithelium (Wang, 2007).
Because RMLC phosphorylation is strongly reduced in rok mutant cells, rok clones were used to examine the function of apical myosin activity. rok mutant follicle cells divide normally, form a monolayered follicular epithelium, and retain polarity. However, rok mutant cells fail to adopt a normal shape. As a consequence, the epithelium is flatter in these regions than it is in regions with rok activity. Optical sections at the level of the zonula adherens show that rok mutant cells are also stretched compared to neighboring wild-type cells. Furthermore, egg chambers with large follicle cell clones develop abnormal shapes as the cyst bulges outwards in the area of the clones. These results show that rok is required for follicle cell and egg-chamber shape, and indicate that the function of the apical myosin activity is to prevent flattening and stretching of epithelial cells (Wang, 2007).
To test the function of the apical myosin activity directly, follicle cell clones were generated using a null mutation for spaghetti squash (sqh). sqh encodes RMLC and was previously shown to be required for other aspects of egg-chamber morphogenesis, like cyst separation and follicle cell migration (Karess, 1991; Edwards, 1996). Follicle cells lacking RMLC activity are extremely flat and appear stretched. In many egg chambers with sqh clones, gaps were found in the follicular epithelium, suggesting that stretching of the follicle cells eventually disrupts the monolayer. The flat sqh mutant cells retain polarity, as revealed by the localization of Discs large (Dlg), a marker for the region where the septate junctions are formed, and the localization of the apical marker aPKC. The change in the shape of the follicle cells is accompanied by a change in the morphology of the egg chamber. Although those regions of the egg chamber covered by wild-type follicle cells retain a normal shape, the germline cyst bulges out in regions covered by sqh mutant cells. In summary, the morphological defects in the sqh clones are very similar to the defects in the rok mutant clones, although the sqh phenotype is much stronger. The stronger morphological defects in sqh mutants are consistent with the finding that RMLC activity is only reduced in rok, whereas it is abolished in sqh mutant cells (Wang, 2007).
sqh function is also required for cytokinesis, and, consistent with this, epithelia with sqh mutant clones show a reduced number of phospho-Histone H3-positive cells, huge nuclei, and abnormally large cells. To examine whether these defects contribute to the morphological defects, epithelia with clones mutant for diaphanous (dia), another gene required for cytokinesis, were examined. Using the weak allele dia5, follicle cell clones were identifed showing cytokinesis defects in the presence of a normal actin cortex. During early oogenesis, these clones retain a rectangular shape, do not flatten, and the underlying cyst bulges out only very mildly. Late clones show no outward bulging over the growing oocyte and maintain a normal cell shape, with the exception that the cells are bigger because of the absence of cytokinesis. Thus, cytokinesis defects alone do not affect the rectangular shape of the follicle cells, and they affect the shape of the egg chamber only mildly and only during early oogenesis. Importantly, the morphological defects are fully penetrant in sqh mutant follicle cell clones. It is therefore concluded that the morphological defects in egg chambers with sqh clones are the result of the loss of apical myosin activity (Wang, 2007).
The epithelial deformations in sqh clones suggest a stress that is acting on the epithelium. The outward bulging of the cyst further suggests that the origin of this stress is the volume increase of the growing cyst. Wild-type cells might resist this stress because of the myosin activity at the apical cortex that is facing the cyst, whereas sqh mutant cells collapse. To test this hypothesis, cyst growth was blocked by using a chromosome carrying an ovoD1 mutation. ovoD1 is a dominant female-sterile mutation that is normally applied in germline mosaics. Importantly, the ovoD1 phenotype is restricted to the germline and does not affect the somatic epithelium. The ovoD1 harbouring chromosome that was used in this experiment leads to a growth arrest after stage 4 resulting in small stage 6 egg chambers, which later degenerate (Wang, 2007).
sqh follicle cell clones were genrated in parallel in wild-type and in ovoD1 mutant backgrounds and cyst and epithelial shape was analyzed. Strikingly, sqh mutant follicle cells maintain their rectangular shape when cyst growth is blocked, whereas sqh cells are deformed when the cyst grows. Moreover, the cyst bulges out underneath the sqh clones only in the wild-type background, but not in the ovoD1 mutant cysts. Thus, myosin activity is required for epithelial and egg-chamber shape only if the cyst is growing. It is therefore concluded that epithelial myosin activity counteracts the force from the growing cyst (Wang, 2007).
How could myosin activity counteract stress from the growing cyst mechanistically? In Dictyostelium, it has been shown that the cell membrane is able to resist deformations induced by a cell poker, revealing stiffness of the cortex. In myosin mutants, the cortical stiffness is greatly reduced, indicating that stiffness is generated by myosin-mediated contractions within the actin cortex. Consistent with this, in vitro studies demonstrated that myosin activity increases the stiffness of crosslinked actin filaments by a factor of 100. Stiffness is generated by diminishing thermal fluctuations within a crosslinked actin network. Myosin is able to suppress these fluctuations by mediating contractions of actin filaments between crosslink points. It is proposed that stiffness is a crucial feature of the apical epithelial cortex in response to the stress emanating from the growing cyst, and that myosin regulates the stiffness by generating tension between actin crosslink points (Wang, 2007).
The pattern of myosin activity reflects the organization of the actin cytoskeleton. The stress fiber-like pattern at the basal cortex reveals activity in the parallel actin arrays, and this activity leads to egg-chamber elongation. In contrast to this polarized pattern of myosin, the apical pattern shows no uniform direction, indicating that actin filaments of all orientations contract. This suggests that the actin filaments at the apical cortex are crosslinked like a net. Thermal fluctuations are higher in actin networks compared to bundled actin filaments. A netlike organization of the actin filaments is therefore consistent with the model, in which myosin-mediated contractions increase cortical stiffness by suppressing thermal fluctuations within the net (Wang, 2007).
The follicular epithelium responds to the cyst growth by increasing the epithelial surface by cell proliferation. The signal that induces mitosis is unknown. These data raise the possibility that the actomyosin cytoskeleton is involved in the coordination of cyst growth and epithelial proliferation. It is likely that the apical cortex, which is stiffened by myosin, perceives the volume increase of the growing cyst as a further tension increase in the crosslinked actin filaments. It is speculated that tension increase above a certain threshold triggers mitosis in the epithelium. The resulting cell divisions lead to an enlarged epithelial surface and thereby to a tension decrease at the apical cortex. The coupling of tension increase and cell proliferation adapts the growth of the epithelium to the volume increase of the cyst and prevents epithelial rupture. The role of tension in regulating cell growth was proposed in the past and has been demonstrated recently in cell culture experiments (Wang, 2007).
If cyst growth and epithelial proliferation are coupled, follicle cell division should be reduced when the cyst volume does not increase. Notably, a dramatic reduction was found in cell division in ovoD1 mutant ovarioles, in which growth is blocked. Consistent with this, it has been reported that block of cyst growth induced by germline clones mutant for the Drosophila Insulin receptor and dMyc does not result in excess follicle cells. These results show that cyst growth and epithelial growth are coupled. However, they allow no conclusion about the coupling mechanism (Wang, 2007).
The restriction of myosin activity to the apical cortex of the epithelium is mediated by at least three different mechanisms. First, myosin phosphorylation at the apical cortex is achieved by apical localization of Rok. Rok is also regulated by the small GTPase Rho1. rho1 mutant follicle cell clones show reduced apical myosin phosphorylation and cell flattening, suggesting that Rho1 binding enables Rok to phosphorylate myosin. In contrast to rok mutant clones, rho1 mutant cells have large nuclei and an increased cell size, indicating that Rho1 is also required for cytokinesis. The second mechanism that restricts myosin activity to the apical cortex is the anchoring of active myosin by the Baz/aPKC/Par-6 complex. The third mechanism is the inhibition of myosin at the lateral and basal cortex via PP1β9C-mediated dephosphorylation. In the future, it will be important to find additional components regulating apical myosin activity, and to find out whether myosin activity is also in other epithelia restricted to certain domains (Wang, 2007).
Subdividing proliferating tissues into compartments is an evolutionarily conserved strategy of animal development. Signals across boundaries between compartments can result in local expression of secreted proteins organizing growth and patterning of tissues. Sharp and straight interfaces between compartments are crucial for stabilizing the position of such organizers and therefore for precise implementation of body plans. Maintaining boundaries in proliferating tissues requires mechanisms to counteract cell rearrangements caused by cell division; however, the nature of such mechanisms remains unclear. This study quantitatively analyzed cell morphology and the response to the laser ablation of cell bonds in the vicinity of the anteroposterior compartment boundary in developing Drosophila wings. Mechanical tension was found to be approximately 2.5-fold increased on cell bonds along this compartment boundary as compared to the remaining tissue. Cell bond tension is decreased in the presence of Y-27632, an inhibitor of Rho-kinase whose main effector is Myosin II. Simulations using a vertex model demonstrate that a 2.5-fold increase in local cell bond tension suffices to guide the rearrangement of cells after cell division to maintain compartment boundaries. These results provide a physical mechanism in which the local increase in Myosin II-dependent cell bond tension directs cell sorting at compartment boundaries (Landsberg, 2009).
A long-standing hypothesis to explain the maintenance of compartment boundaries is based on differential cell adhesion (or cell affinity). Cell adhesion molecules required for the maintenance of compartment boundaries, however, have not been identified. More recently, it has been proposed that actin-myosin-based tension is important for keeping the dorsoventral compartment boundary of the developing Drosophila wing smooth and straight. However, whether a similar mechanism operates at the anteroposterior compartment boundary (A/P boundary) is unclear. Moreover, a physical measurement of differential mechanical tension at compartment boundaries has not been reported. Furthermore, whether and how differential mechanical tension governs cell sorting at compartment boundaries is not well understood (Landsberg, 2009).
To test whether actin-myosin-based tension is increased at the A/P boundary, the levels of Filamentous (F)-actin and nonmuscle Myosin II (Myosin II) were quantified. The A/P boundary in the wing disc epithelium was particularly well defined by the cell bonds located at the level of adherens junctions, indicating that mechanisms maintaining the boundary operate at this cellular level. F-actin and the regulatory light chain of Myosin II (encoded by spaghetti squash, sqh) were increased at these cell bonds along the A/P boundary. Cell bonds displaying elevated levels of Myosin II correlate with decreased levels of Par3 (Bazooka in Drosophila), a protein organizing cortical domains, at the dorsoventral compartment boundary and during germ-band extension in Drosophila embryos. Likewise, Bazooka was decreased at cell bonds along the A/P boundary, indicating a common mechanism of complementary protein distribution of Myosin II and Bazooka. The level of E-cadherin, a component of adherens junctions, was not altered along the A/P boundary (Landsberg, 2009).
To identify signatures of increased tension in the vicinity of the A/P boundary, the morphology of cells were quantitatively analyzed at the level of adherens junctions. Line tension and mechanical properties of cells have been proposed to contribute to cell shape and to influence angles between cell bonds. Line tension associated with adherens junctions, here termed cell bond tension, can be defined as the work, per unit length, performed as a cell bond changes its length. Cell bond tension results from actin-myosin bundles and other structural components at junctional contacts that generate tensile stresses. Wing discs from late-third-instar larvae were stained for E-cadherin and engrailed-lacZ, a marker for the posterior compartment. Cell bonds were identified, and morphological parameters were analyzed. Adjacent anterior and posterior cells (A1 and P1, respectively) displayed a significantly enlarged apical cross-section area compared to cells farther away from the compartment boundary, indicating that apposition of anterior and posterior cells alters specifically the properties of A1 and P1 cells. Angles between adjacent cell bonds along the A/P boundary were larger compared to angles between bonds of the remaining cells and were significantly smaller in mutants for Myosin II heavy chain (encoded by zipper; zip2/zipEbr). Thus, the unique morphology of A1 and P1 cells depends on Myosin II. These data are consistent with an increased Myosin II-based tension of cell bonds located along the A/P boundary (Landsberg, 2009).
Cells on opposite sides of the A/P boundary differ in gene expression. The homeodomain-containing proteins Engrailed and Invected as well as the Hedgehog ligand are only expressed on the posterior side. The Hedgehog signal is transduced exclusively on the anterior side. Hedgehog signal transduction and the presence of Engrailed and Invected are required to maintain this compartment boundary. Whether the altered cell morphology at the A/P boundary could be reproduced by ectopically juxtaposing Hedgehog signaling and non-Hedgehog signaling cells was tested. Clones of cells that expressed Hedgehog from a transgene and that were also mutant for the gene smoothened (encoding an essential transducer of the Hedgehog pathway) were generated. In the P compartment, which is refractory to Hedgehog signal transduction, clones displayed a normal morphology. In the A compartment, a response to Hedgehog that is secreted by the clones is elicited in the surrounding wild-type cells. These clones had a rounder appearance, and at the clone border, but not away from it, apical cross-section area and bond angles were increased. Similarly, juxtaposing cells expressing engrailed and invected with cells that are mutant for these genes resulted in increased apical cross-section area and increased bond angles at the clone border. It is concluded that the morphology that is characteristic of cells at the A/P boundary can be imposed on cells within a compartment by juxtapositioning cells with different activities of Hedgehog signal transduction or Engrailed and Invected (Landsberg, 2009).
Ablating cell bonds generates cell vertex displacements, providing direct evidence for tension on cell bonds. Individual cell bonds were ablated by using a UV laser beam focused in the plane of the adherens junctions. Single-cell bonds were cut, and the displacement of vertices of neighboring cells, visualized by E-cadherin-GFP, was recorded. The P compartment was visualized by expression of GFP-gpi under control of the engrailed gene via the GAL4/UAS system. The increase in distance between the two vertices of the ablated cell bond and the initial velocity of this vertex separation were analyzed. The ratio of initial velocities in response to cell bond ablation is a measure of the tension ratio on these cell bonds. Initial velocity and extent of vertex separation were indistinguishable between anterior (A/A) and posterior (P/P) cell bonds located away from the A/P boundary. This was also the case when specifically cell bonds between the first and second row of anterior cells were ablated. By contrast, ablation of bonds between adjacent anterior and posterior cells (A/P cell bonds) gave rise to a larger vertex separation. This result was not due to the fact that A/P cell bonds have a preferred orientation. Moreover, the initial velocity of ablated A/P bonds was 2.37-fold higher compared to the mean of initial velocities of A/A and P/P bonds. This value provides an estimate of the ratio λ of cell bond tension along the A/P boundary relative to the average tension of cell bonds. In the presence of the Rho-kinase inhibitor Y-27632, the ratio of initial velocity of vertex separation of A/P cell bonds relative to A/A cell bonds was reduced to 1.46. Given that Myosin II is the main effector of Rho-kinase, these results strongly suggest that Myosin II-based tension acting on cell bonds is locally increased along the A/P boundary (Landsberg, 2009).
To quantify λ by an independent method, the displacement field was calculated after laser ablation. Using a vertex model, two populations of adjacent cells were introduced and cell bond ablations were simulated, varying λ between 1 and 4. When λ = 2.5, the vertex displacement, and in particular the anisotropy of displacements, in the simulations closely matched the vertex displacements in the experiment. In the vertex model, λ = 2.5 also resulted in increased bond angles at the interface of the two cell groups, similar to the A/P boundary in the wing disc. Thus, on the basis of two different methods, the data demonstrate that cell bond tension is increased approximately 2.5-fold along the A/P boundary compared to the remaining tissue (Landsberg, 2009).
To test whether a 2.5-fold increase in cell bond tension is sufficient to maintain a compartment boundary, the vertex model was used to simulate the growth of two adjacent cell populations for λ = 1, 2.5, and 4. For λ = 1, the interface between two growing cell populations became increasingly irregular. By contrast, for λ = 2.5 and 4, a well-defined interface was maintained. Moreover, corresponding changes in cell bond tension at borders of simulated clones resulted in the morphology and sorting behavior of cell patches that resembled those of experimental cell clones compromised for Hedgehog signal transduction or Engrailed and Invected activity. The roughness of the interface in the simulations decreased with increasing λ, showing that cell bond tension is sufficient to maintain straight interfaces between growing cell populations. For λ = 2.5, the roughness of the interface was still larger than the roughness of the A/P boundary in wing discs. This suggests that additional mechanisms might contribute to further reduce the roughness of the A/P boundary. Also, because of the uncertainty of the mechanical properties of A1 and P1 cells, which differ from those of the remaining cells, the value of λ, inferred from laser ablation of cell bonds, might be underestimated. Remarkably, the roughness of the A/P boundary could be altered in mutant conditions. In zip2/zipEbr mutant wing discs, the roughness of the compartment boundary was significantly larger than in controls, demonstrating a role for Myosin II in maintaining a sharp and straight A/P boundary (Landsberg, 2009).
In summary, by applying physical approaches and quantitative imaging, this work for the first time demonstrates and quantifies an increase in tension confined to the cell bonds along the A/P boundary. Moreover, simulations show that this increase in tension suffices to maintain a stable interface between two proliferating cell populations. Genetic studies demonstrated that cells of the two compartments differ in their expression profiles and signaling activities. It has therefore been proposed that biophysical properties of cells within the P compartment differ from those within the A compartment, and that such differences could drive cell sorting. When quantifying cell morphology and vertex displacements after laser ablation, no differences were detected in the biophysical properties of cells between the two compartments. However, the two rows of abutting A and P cells show clear differences in biophysical properties from other cells. Most importantly, the cell bond tension along the A/P boundary is increased. Cell divisions in the vicinity of the A/P boundary were randomly oriented in the epithelial plane. Thus, taken together with the simulations, these results suggest a sorting mechanism by which an increased cell bond tension guides the rearrangement of cells after cell division to maintain a straight interface. Increased cell bond tension and the roughness of the A/P boundary depend on Rho kinase activity and Myosin II, indicating a role for actin-myosin-based tension in this process. Because cell bond tension also depends on cell-cell adhesion, differences in the adhesion between A1 and P1 cells as compared to the remaining cells might also contribute to sorting. The heterotypic, but not homotypic, interaction of molecules presented on the surface of A and P cells might trigger the local increase in cell bond tension. Hedgehog signal transduction and the presence of Engrailed and Invected might control the expression of these heterotypically interacting molecules. These data indicate an important role for cell bond tension directing cell sorting during animal development (Landsberg, 2009).
Cell rearrangements shape the Drosophila embryo via spatially regulated changes in cell shape and adhesion. This study of axis elongation (germband extension) shows that Bazooka/Par-3 (Baz) is required for the planar polarized distribution of myosin II and adherens junction proteins and polarized intercalary behavior is disrupted in baz mutants. The myosin II activator Rho-kinase is asymmetrically enriched at the anterior and posterior borders of intercalating cells in a pattern complementary to Baz. Loss of Rho-kinase results in expansion of the Baz domain, and activated Rho-kinase is sufficient to exclude Baz from the cortex. The planar polarized distribution of Baz requires its C-terminal domain. Rho-kinase can phosphorylate this domain and inhibit its interaction with phosphoinositide membrane lipids, suggesting a mechanism by which Rho-kinase could regulate Baz association with the cell cortex. These results demonstrate that Rho-kinase plays an instructive role in planar polarity by targeting Baz/Par-3 and myosin II to complementary cortical domains (de Matos Simões, 2010).
The spatially regulated activity of protein kinases with multiple substrates provides an efficient strategy for the control of cell polarity in different contexts. This study shows that Rho-kinase is an asymmetrically localized protein that plays an instructive role in planar polarity in the Drosophila embryo by excluding its substrate Baz/Par-3 from the cell cortex. Rho-kinase prevents expansion of the Baz domain and Baz in turn directs the localization of contractile and adherens junction proteins that are required for axis elongation, converting a localized source of kinase activity into a robust bias in polarized cell behavior. The effect of Rho-kinase on Baz planar polarity appears to be independent of its role in regulating myosin II, as Baz localization is not affected in myosin mutants and activated myosin does not reproduce the effects of Rho-kinase in culture. Instead, Rho-kinase can directly phosphorylate the Baz C-terminal coiled-coil domain that is required for Baz association with the cortex. Deletions within the Baz C-terminal domain or replacement of the Baz C-terminus with a heterologous phospholipid binding motif abolish Baz planar polarity in vivo. These results are consistent with a model in which Rho-kinase directly inhibits the association of the Baz C-terminal domain with specific regions of the cell cortex (de Matos Simões, 2010).
Rho-kinase has been shown to phosphorylate mammalian Par-3 in cultured cells, disrupting its interaction with the Par complex proteins Par-6 and aPKC (Nakayama, 2008). The Par complex is necessary for some aspects of epithelial organization but dispensable for others. Par-6 and aPKC are not required for Baz planar polarity in Drosophila, suggesting that the role of Rho-kinase in this process is unlikely to occur through a similar mechanism. This study provides evidence for a different mechanism of regulation by Rho-kinase involving the Baz C-terminal domain, which is phosphorylated by Rho-kinase in vitro and is necessary for Baz planar polarity in vivo. The Baz C-terminus has been shown to bind directly to phosphoinositide membrane lipids including PI(3,4,5)P3, PI(3,4)P2 and PIP (Krahn, 2010). This study shows that Rho-kinase inhibits the association of Baz with phosphoinositide membrane lipids in vitro, consistent with a model in which Rho-kinase directly regulates Baz association with the cortex. Alternatively, Rho-kinase could regulate Baz localization indirectly through other proteins that interact with the Baz C-terminal domain. Despite potential differences in the mechanism, these results demonstrate that the regulation of Par-3 localization or activity by Rho-kinase is a conserved feature of cell polarity in Drosophila and mammals (de Matos Simões, 2010).
The results demonstrate that Rho-kinase is an asymmetrically localized protein that initiates a cascade of events required for the planar polarized distribution of contractile and adherens junction proteins in intercalating cells. The upstream signals that generate localized Rho-kinase activity are not known. Differences between cells conferred by striped or graded patterns of gene expression orient cell movement during axis elongation, and AP patterning genes expressed in stripes are necessary for the asymmetric localization of Rho-kinase. These findings raise the possibility that planar cell polarity may be generated by the local activation of a Rho GTPase signaling pathway. The Drosophila genome contains 21 RhoGEFs and 19 RhoGAPs that are candidate upstream regulators in this process. Rho GTPase pathways are activated by a number of upstream signals including G protein-coupled receptors, receptor tyrosine kinases, cytokine receptors, and cell-cell and cell-substrate adhesion. Identification of the signals upstream of Rho-kinase will help to elucidate the spatial cues that initiate planar polarity in the Drosophila embryo (de Matos Simões, 2010).
The role of Rho-kinase in planar cell polarity is reinforced by the effect of Baz on the localization of contractile and adherens junction proteins. The relationship between Baz and myosin II is complex. In the C. elegans zygote, a contractile myosin network carries PAR-3 to the anterior cell cortex, suggesting a positive relationship between these proteins. In other cell types myosin appears to be dispensable for Baz localization. PAR-3 is required to sustain myosin contractility in C. elegans and Drosophila, and Baz promotes myosin apical localization during C. elegans gastrulation and in the Drosophila follicular epithelium. The ectopic association of myosin with DV cell boundaries in baz mutants, and the complementary distributions of Baz and myosin in several contexts, raise the possibility of inhibitory effects of Baz on myosin. This regulation could also occur indirectly through effects of Baz on apical-basal polarity (de Matos Simões, 2010).
Differential adhesion is sufficient to drive cell sorting in culture and has been proposed to influence tissue morphogenesis in vivo. This study shows that Rho-kinase and Baz regulate the planar polarized localization of the adherens junction protein β-catenin. Rho-kinase has been shown to downregulate adhesion in culture, an activity that is thought to occur through myosin II, which can play positive and negative roles in junctional stabilization. The ability of Rho-kinase to exclude the Baz/Par-3 junctional regulator from the cortex suggests an alternative mechanism for the regulation of adherens junctions by Rho GTPases. These results suggest that Rho-kinase can both promote contractility and inhibit adhesion, providing a single molecular mechanism linking cortical contraction with adherens junction disassembly during tissue morphogenesis (de Matos Simões, 2010).
Morphogenesis of epithelial tissues relies on the interplay between cell division, differentiation and regulated changes in cell shape, intercalation and sorting. These processes are often studied individually in relatively simple epithelia that lack the complexity found during organogenesis when these processes might all coexist simultaneously. To address this issue, this study makes use of the developing fly retinal neuroepithelium. Retinal morphogenesis relies on a coordinated sequence of interdependent morphogenetic events that includes apical cell constriction, localized alignment of groups of cells and ommatidia morphogenesis coupled to neurogenesis. Live imaging was used to document the sequence of adherens junction (AJ) remodelling events required to generate the fly ommatidium. In this context, it was demonstrated that the kinases Rok and Drak function redundantly during Myosin II-dependent cell constriction, subsequent multicellular alignment and AJ remodelling. In addition, it was shown that early multicellular patterning characterized by cell alignment is promoted by the conserved transcription factor Atonal (Ato). Further ommatidium patterning requires the epidermal growth factor receptor (EGFR) signalling pathway, which transcriptionally governs Rho-kinase (rok) and Death-associated protein kinase related (Drak)-dependent AJ remodelling while also promoting neurogenesis. In conclusion, this work reveals an important role for Drak in regulating AJ remodelling during retinal morphogenesis. It also sheds new light on the interplay between Ato, EGFR-dependent transcription and AJ remodelling in a system in which neurogenesis is coupled with cell shape changes and regulated steps of cell intercalation (Robertson, 2013).
In Drosophila, Rok seems to be the main kinase responsible for phosphorylating the Myosin regulatory light chain (Sqh) during epithelial patterning and apical cell constriction. This is the case for the activation of MyoII during intercalation as germband extension proceeds, but also during various instances of compartment boundary formation and cell sorting situations in the embryo and in the wing imaginal disc. The current work reveals that in the constricting cells of the MF, Rok functions redundantly with Drak, a kinase recently shown to phosphorylate Sqh both in vitro and in vivo (Neubueser, 2010). It is noteworthy that previous work has shown that RhoGEF2 is not required for cell constriction in the MF, suggesting that perhaps another guanine exchange factor (GEF) might function redundantly with RhoGEF2 to promote cell constriction. These data on Drak reinforce the idea that redundancies exist in this context. Because the RhoA (Rho1 -- FlyBase) loss of function abolishes this cell response entirely, it would be expected that Drak function is regulated by RhoA. In addition, the current data indicate that Drak acts redundantly with Rok during MyoII-dependent multicellular alignment and AJ remodelling during ommatidia patterning. It will be interesting to test whether Drak functions in other instances of epithelial cell constriction or MyoII-dependent steps of AJ remodelling in other developmental contexts in Drosophila (Robertson, 2013).
This study demonstrates a two-tiered mechanism regulating the planar polarization of MyoII and Baz. In the constricting cells in the posterior compartment, MyoII and Baz are segregated from one another and this is exacerbated by the wave of cell constriction in the MF. Upon Ato-dependent transcription in the MF cells, this segregated pattern of expression is harnessed and these factors become planar polarized at the posterior margin of the MF. This is independent of the core planar polarity pathway including the Fz receptor and is accompanied by a striking step of multicellular alignment. Previous work has demonstrated that Ato upregulates E-Cad transcription at the posterior boundary of the MF. In addition, apical constriction leads to an increase in E-Cad density at the ZA. The current data are therefore consistent with both hh-dependent constriction and ato-dependent transcriptional upregulation of E-Cad promoting differential adhesion, thus leading to a situation in which the ato+ cells maximize AJ contacts between themselves and minimize contact with the flanking cells that express much less E-Cad at their ZA. This typically leads to a preferential accumulation of cortical MyoII at the corresponding interface. Such actomyosin cables are correlated with increased interfacial tension, and it is proposed that this is in turn responsible for promoting cell alignment. Unfortunately, the very small diameter of these constricted cells precludes direct measurements of the AJ-associated tension using laser ablation experiments (Robertson, 2013).
Supra-cellular cables of MyoII have been previously associated with cell alignment in various epithelia and have also been observed at the boundary of sorted clones, whereby cells align at a MyoII-enriched interface. Interestingly, this study found that the actomyosin cable defining the posterior boundary of the MF is also preferentially enriched for Rok, a component of the T1, MyoII-positive AJ in the ventral epidermis (Simoes Sde, 2010). This indicates an important commonality between actomyosin cable formation during cell sorting and the process of cell intercalation. However, unlike during intercalation, this study found that in the developing retina baz is largely dispensable for directing the pattern of E-Cad and actomyosin planar polarization. Further work will therefore be required to understand better the relationship between Baz and E-Cad at the ZA during ommatidia morphogenesis. It is speculated that the creation of a high E-Cad versus low E-Cad boundary in the wake of the MF might be sufficient to promote Rok and MyoII enrichment at the posterior AJs. This posterior Rok and MyoII enrichment might perhaps prevent E-Cad accumulation by promoting E-Cad endocytosis, as has been recently shown in the fly embryo (Robertson, 2013).
This study has used live imaging to define a conserved step of ommatidia patterning that consists of the coalescence of the ommatidial cells' AJs into a central vertex to form a 6-cell rosette. The corresponding steps of AJ remodelling require Rok, Drak, Baz and MyoII, a situation compatible with mechanisms previously identified during cell intercalation in the developing fly embryo. The steps of AJ remodelling required to transform lines of cells into 5-cell pre-clusters are transcriptionally regulated downstream of EGFR in a ligand-dependent manner. Interestingly, in the eye EGFR signalling is activated in the cells that form lines and type1-arcs in the wake of the MF and, thus, are undergoing AJ remodelling. Previous work examining tracheal morphogenesis in the fly has demonstrated that interfaces between cells with low levels versus high levels of EGFR signalling correlate with MyoII-dependent AJ remodelling in the tracheal placode. This situation resembles that which is described in this study in the wake of the MF. In the eye, however, it was found that EGFR signalling is not required to initiate cell alignment. Nevertheless, taken together with work in the tracheal placode and previous studies related to multicellular patterning in the developing eye, this work indicates a conserved function for the EGFR signalling pathway in promoting MyoII-dependent AJ remodelling. This leaves open several interesting questions; for example, it is not presently clear how EGFR signalling can promote discrete AJ suppression and elongation. It is, however, tempting to speculate that previously described links between EGFR signalling and the expression of E-Cad or Rho1 might play a role during this process (Robertson, 2013).
The rok gene maps to chromosome region 15A1 on the X chromosome. To generate loss-of-function mutations in rok, the assumption was made that hemizygous rok mutant males would be lethal, but that such lethality could be rescued by either a genomic duplication that covers the region [Tp(1; 2)r+75c] or by a rok cDNA expressed under the control of the ubiquitously active tubulin-1alpha promoter (tubP-Drok). Screening EMS mutagenized flies led to the identification of two X-linked lethal mutants that were rescued by Tp(1; 2)r+75c and tubP-Drok, and failed to complement each other. These were named Drok1 and Drok2 (Winter, 2001).
To determine the molecular nature of the rok mutations, genomic sequencing of the mutant DNA was performed. The coding sequence of rok is composed of 10 exons. A single nucleotide transition of G to A was found in the Drok2 mutant at the 3' end of intron 1, changing the conserved AG at the splice acceptor site to AA. Quantitative RT-PCR revealed no change in the transcript level in Drok2 hemizygous larvae compared to control. However, sequence analysis of the Drok2 mutant cDNA revealed that the transcript has a single base deletion of guanine at the beginning of exon 2, due to the use of a splice acceptor site one nucleotide 3' of the original site. The mutant mRNA encodes only the first 21 amino acids (aa) of Rok, followed by a 35 aa random peptide and a stop codon, before the start of the kinase domain. Hence, Drok2 is likely to be a strong loss-of-function, if not a null mutation. Since the molecular nature of the Drok1 mutation has not been determined, the Drok2 allele was used for the rest of this study (Winter, 2001).
The global cell movements that shape an embryo are driven by intricate changes to the cytoarchitecture of individual cells. In a developing embryo, these changes are controlled by patterning genes that confer cell identity. However, little is known about how patterning genes influence cytoarchitecture to drive changes in cell shape. This paper analyzes the function of the folded gastrulation gene (fog), a known target of the patterning gene twist. Analysis of fog function therefore illuminates a molecular pathway spanning all the way from patterning gene to physical change in cell shape. Secretion of Fog protein is apically polarized, making this the earliest polarized component of a pathway that ultimately drives myosin to the apical side of the cell. fog is both necessary and sufficient to drive apical myosin localization through a mechanism involving activation of myosin contractility with actin. This contractility driven form of localization involves RhoGEF2 and the downstream effector Rho kinase. This distinguishes apical myosin localization from basal myosin localization; the latter does not require actinomyosin contractility or FOG/RhoGEF2/Rho-kinase signaling. Furthermore, once localized apically, myosin continues to contract. The force generated by continued myosin contraction is translated into a flattening and constriction of the cell surface through a tethering of the actinomyosin cytoskeleton to the apical adherens junctions. Therefore, this analysis of fog function provides a direct link from patterning to cell shape change (Dawes-Hoang, 2005).
The components acting downstream of fog to mediate its effects on the cytoskeleton are largely unknown. One candidate, RhoGEF2 (a guanine nucleotide exchange factor that promotes Rho activation) has been shown to be required for ventral furrow formation and can genetically interact with a fog transgene. However, embryos mutant for RhoGEF2 have a much more severe disruption of ventral furrow formation than embryos mutant for fog and the point at which the products of these two genes interact on a mechanistic or subcellular level is unknown. Recent studies have shown a requirement for RhoGEF2 in controlling actin dynamics/stability during cellularization and have also shown a disruption to myosin localization at gastrulation. Therefore the re-localization of myosin during cellularization and gastrulation was analyzed in RhoGEF2 mutants and previous studies were extended by looking at a potential downstream effector of RhoGEF2 signaling (Dawes-Hoang, 2005).
Embryos mutant for RhoGEF2 localize myosin normally to the forming cellularization front. However, unlike fog mutants, the RhoGEF2 embryos show defects in cellularization, including an irregular, wavy cellularization front. This implies that although RhoGEF2 function is not required to localize myosin to the cellularization front it is required to maintain the normal structure of the cellularization front and that the presence of myosin is not itself sufficient to maintain a straight cellularization front. This is consistent with previous studies of RhoGEF2 mutants and a potential role in controlling actin but not myosin dynamics (Grosshans, 2005; Padash Barmchi, 2005; Dawes-Hoang, 2005 and references therein).
However, despite the defects during early cellularization, RhoGEF2 mutant embryos that reach the end of cellularization look remarkably normal. The irregularity of the cellularization front recovers, particularly in the ventral cells, and both the increased cell depth and basal loss of myosin occur normally in these cells. However, in RhoGEF2 embryos, precisely staged for the onset of gastrulation, there is an absolute failure to re-localize myosin to the apical side of the ventral cells, despite a normal loss of myosin from the basal side of these cells. This is consistent with independent mechanisms controlling the basal loss and apical accumulation of myosin during gastrulation and demonstrates an absolute requirement for RhoGEF2 in apical myosin localization. It also confirms the previous report of RhoGEF2 being required for apical myosin in ventral furrow cells (Nikolaidou, 1994; Dawes-Hoang, 2005).
RhoGEF2 interacts with myosin in other systems through the Rho-kinase family of Ser/Thr kinases that inhibit myosin phosphatase and also directly phosphorylate myosin. Both these activities led to activation of actin binding by myosin and increased actomyosin based contractility. Additional myosin activators include MLCK and citron kinase but the extent to which these different activators play specific or overlapping roles with Rho-kinase is unclear, and the role of any of these myosin activators during Drosophila gastrulation is not known (Dawes-Hoang, 2005).
Therefore embryos were produced mutant for Drosophila Rho-kinase (Drok) by making germline clones of two Drok alleles, both of which produced similar phenotypes. Myosin localizes to the cellularization front of Drok mutant embryos but often does so unevenly and, as for RhoGEF2, the cellularization front is 'wavy'. Unlike the RhoGEF2 mutant embryos, the nuclei of Drok mutant embryos have striking defects, including displacement into the interior of the embryo leaving reduced numbers at the cortex: these remaining nuclei are often of increased size and irregular morphology. It is unclear to what extent these nuclear phenotypes may represent an earlier defect during cell-cycle/nuclear division (Dawes-Hoang, 2005).
Despite these defects, many Drok mutant embryos complete cellularization and though the increased depth of cellularization in ventral cells is difficult to discern, basal loss of myosin proceeds normally. However, Drok mutant embryos show a complete failure to localize myosin to the apical side of the ventral cells at the onset of gastrulation. At later stages of gastrulation, the outer layer of wild-type embryos consists of a single cell layered epithelium that folds in specific locations during germband extension. In Drok mutant embryos this morphology is severely disrupted and the outer epithelium becomes multilayered and irregular, containing large often rounded cells. Drok is therefore required to maintain epithelial integrity (Dawes-Hoang, 2005).
Both Drok and RhoGEF2 mutant embryos show defects during cellularization and then fail to localize myosin to the apical side of ventral cells at gastrulation. However, it is unlikely that the earlier cellularization defects are what prevent the later apical myosin localization because many other cellularization mutants, such as nullo, display severe cellularization defects but still go on to localize myosin to the apical side of ventral cells at the onset of gastrulation. The failure of Drok and RhoGEF2 mutant embryos to localize myosin apically during gastrulation therefore probably reflects a direct requirement for both these genes in the apical localization of myosin. Despite these disruptions to gastrulation, Drok embryos do still produce Fog protein that is as punctate and apically concentrated as in wild-type embryos. This is therefore consistent with a model whereby Drok driven activation of myosin contractility drives myosin apically in response to fog and RhoGEF2 signaling (Dawes-Hoang, 2005).
Taken together these data suggest the following model. Expression of the patterning gene twi in the prospective mesoderm cells results in activation of fog transcription. The resulting Fog protein is then secreted from the apical surface of the cells and this signal activates fog receptors. The degree to which this activation is paracrine versus autocrine has yet to be determined. The apically activated receptors trigger a transduction pathway involving the G-alpha subunit, Concertina, and the Rho activator RhoGEF2. A downstream target of this pathway is Rho-kinase, which in turn activates the ability of myosin to interact and contract with actin in this sub-apical region of the cell. A localized source of activated actin-myosin contractility initiates an active motor-driven mechanism of myosin localization that concentrates contractile myosin to the apical side of the cell. This actin-myosin network is tethered to the cell surface through adherens junctions. Contraction of this network therefore puts tension on the junctions, pulling them into a completely apical location and flattening the domed apical surface in the process. Continued contraction exerts further tension and ultimately pulls the junctions together so much that the entire apical cell surface constricts. Intriguingly, RhoGEF2 protein can associate with the tips of microtubules in cultured cells. The extent to which this may add to a polarization of the fog pathway during gastrulation and how this ties in with the above model will therefore be interesting avenues for further investigation. It will also be important to examine any changes to the actin and microtubule organization of these cells (Dawes-Hoang, 2005).
Spatiotemporally regulated actomyosin contractility generates the forces that drive epithelial cell rearrangements and tissue remodeling. Phosphorylation of the myosin II regulatory light chain (RLC) promotes the assembly of myosin monomers into active contractile filaments and is an essential mechanism regulating the level of myosin activity. However, the effects of phosphorylation on myosin localization, dynamics, and function during epithelial remodeling are not well understood. In Drosophila, planar polarized myosin contractility is required for oriented cell rearrangements during elongation of the body axis. This study shows that regulated myosin phosphorylation influences spatial and temporal properties of contractile behavior at molecular, cellular, and tissue length scales. Expression of myosin RLC variants that prevent or mimic phosphorylation both disrupt axis elongation, but have distinct effects at the molecular and cellular levels. Unphosphorylatable RLC produces fewer, slower cell rearrangements, whereas phosphomimetic RLC accelerates rearrangement and promotes higher-order cell interactions. Quantitative live imaging and biophysical approaches reveal that both phosphovariants reduce myosin planar polarity and mechanical anisotropy, altering the orientation of cell rearrangements during axis elongation. Moreover, the localized myosin activator Rho-kinase is required for spatially regulated myosin activity, even when the requirement for phosphorylation is bypassed by the expression of phosphomimetic myosin RLC. These results indicate that myosin phosphorylation influences both the level and the spatiotemporal regulation of myosin activity, linking molecular properties of myosin activity to tissue morphogenesis (Kasza, 2014).
Altun-Gultekin, Z. F., et al. (1998). Activation of Rho-dependent cell spreading and focal adhesion biogenesis by the v-Crk adaptor protein. Mol. Cell. Biol. 18(5): 3044-3058. PubMed Citation: 9566923
Amano, M., et al. (1996). Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase) J. Biol. Chem. 271: 20246-20249. 8702756
Amano M., Chihara K., Kimura K., Fukata Y., Nakamura N., Matsuura Y. and Kaibuchi K. (1997). Formation of actin stress fibers and focal adhesions enhanced by Rho-kinase. Science, 275: 1308-1311. 9036856
Amano, M., et al. (1999). The COOH terminus of Rho-kinase negatively regulates rho-kinase activity. J. Biol. Chem. 274(45): 32418-24. 10542285
Amano M., Fukata Y. and Kaibuchi K. (2000). Regulation and functions of rho-associated kinase. Exp. Cell Res. 261: 44-51. 11082274
Arimura, N., et al. (2000). Phosphorylation of collapsin response mediator protein-2 by Rho-kinase. Evidence for two separate signaling pathways for growth cone collapse. J. Biol. Chem. 275(31): 23973-80. 10818093
Barros, C. S., Phelps, C. B. and Brand, A. H. (2003). Drosophila nonmuscle myosin II promotes the asymmetric segregation of cell fate determinants by cortical exclusion rather than active transport. Dev. Cell 5: 829-840. 14667406
Betschinger, J., Mechtler, K., and Knoblich, J.A. (2003). The Par complex directs asymmetric cell division by phosphorylating the cytoskeletal protein Lgl. Nature 422: 326-330. 12629552
Billuart, P., Winter, C. G., Maresh, A., Zhao, X. and Luo, L. (2001). Regulating axon branch stability. the role of p190 RhoGAP in repressing a retraction signaling pathway. Cell 107(2): 195-207. 11672527
Bito, H., et al. (2000). A critical role for a Rho-associated kinase, p160ROCK, in determining axon outgrowth in mammalian CNS neurons. Neuron 26: 431-441. PubMed Citation: 10839361
Causeret, F., et al. (2004). Distinct roles of Rac1/Cdc42 and Rho/Rock for axon outgrowth and nucleokinesis of precerebellar neurons toward netrin 1. Development 131: 2841-2852. 15151987
Corrigall, D., Walther, R. F., Rodriguez, L., Fichelson, P., Pichaud, F. (2007). Hedgehog signaling is a principal inducer of Myosin-II-driven cell ingression in Drosophila epithelia. Dev Cell 13: 730-742. PubMed ID: 17981140
de Matos Simões, S., et al. (2010). Rho-kinase directs Bazooka/Par-3 planar polarity during Drosophila axis elongation. Dev. Cell 19(3): 377-88. PubMed Citation: 20833361
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(18): 4165-78. 16123312
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. 102(38): 13473-8. 16174742
Diogon, M., et al. (2007). The RhoGAP RGA-2 and LET-502/ROCK achieve a balance of actomyosin-dependent forces in C. elegans epidermis to control morphogenesis. Development 134: 2469-2479. Medline abstract: 17537791
Fagan, J. K., Dollar, G., Lu, Q., Barnett, A., Pechuan Jorge, J., Schlosser, A., Pfleger, C., Adler, P. and Jenny, A. (2014). Combover/CG10732, a novel PCP effector for Drosophila wing hair formation. PLoS One 9: e107311. PubMed ID: 25207969
Gally, C., et al. (2009). Myosin II regulation during C. elegans embryonic elongation: LET-502/ROCK, MRCK-1 and PAK-1, three kinases with different roles. Development 136(18): 3109-19. PubMed Citation: 19675126
Goto, H., et al. (1998). Phosphorylation of vimentin by Rho-associated kinase at a unique amino-terminal site that is specifically phosphorylated during cytokinesis. J. Biol. Chem. 273(19): 11728-36. 9565595
Grosshans, J., Wenzl, C., Herz, H. M., Bartoszewski, S., Schnorrer, F., Vogt, N., Schwarz, H. and Muller, H. A. (2005). RhoGEF2 and the formin Dia control the formation of the furrow canal by directed actin assembly during Drosophila cellularisation. Development 132: 1009-1020. 15689371
Iftinca, M., et al. (2007). Regulation of T-type calcium channels by Rho-associated kinase. Nat Neurosci. 10(7): 854-60. PubMed citation: 17558400
Ishizaki, T, et al. (1996). The small GTP-binding protein Rho binds to and activates a 160 kDa Ser/Thr protein kinase homologous to myotonic dystrophy kinase. EMBO J. 15(8): 1885-1893. PubMed Citation: 8617235
Ishizaki, T., et al. (1997). p160ROCK, a Rho-associated coiled-coil forming protein kinase, works downstream of Rho and induces focal adhesions. FEBS Lett. 404(2-3): 118-124. PubMed Citation: 9119047
Kasza, K. E., Farrell, D. L. and Zallen, J. A. (2014). Spatiotemporal control of epithelial remodeling by regulated myosin phosphorylation. Proc Natl Acad Sci U S A 111(32):11732-7. PubMed ID: 25071215
Kawano, Y., et al. (1999). Phosphorylation of myosin-binding subunit (MBS) of myosin phosphatase by Rho-kinase in vivo. J. Cell Biol. 147(5): 1023-38. 10579722
Kimura, K., et al. (1996). Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273: 245-248. 8662509
Kolega, J. (2003). Asymmetric distribution of myosin IIB in migrating endothelial cells is regulated by a rho-dependent kinase and contributes to tail retraction. Mol. Biol. Cell 14: 4745-4757. 12960430
Kimura, K., et al. (1998). Regulation of the association of adducin with actin filaments by Rho-associated kinase (Rho-kinase) and myosin phosphatase. J. Biol. Chem. 273: 5542-5548. Medline abstract: 9488679
Krahn, M. P., Klopfenstein, D. R., Fischer, N. and Wodarz, A. (2010). Membrane targeting of Bazooka/PAR-3 is mediated by direct binding to phosphoinositide lipids. Curr. Biol. 20: 1-7. PubMed Citation: 21055941
Lammel, U., Bechtold, M., Risse, B., Berh, D., Fleige, A., Bunse, I., Jiang, X., Klambt, C. and Bogdan, S. (2014). The Drosophila FHOD1-like formin Knittrig acts through Rok to promote stress fiber formation and directed macrophage migration during the cellular immune response. Development 141: 1366-1380. PubMed ID: 24553290
Landsberg, K. P., Farhadifar, R., Ranft, J., Umetsu, D., Widmann, T. J., Bittig, T., Said, A., Julicher, F. and Dahmann, C. (2009). Increased cell bond tension governs cell sorting at the Drosophila anteroposterior compartment boundary. Curr Biol 19: 1950-1955. PubMed ID: 19879142
Lee, A., 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. PubMed ID: 15075368
Lee, S. and Helfman, D. M. (2003). Cytoplasmic p21Cip1 is involved in Ras-induced inhibition of the ROCK/LIMK/Cofilin pathway. J. Biol. Chem. 279: 1885-1891. 14559914
Leung, T., et al. (1996). The p160 Rho-binding kinase ROKa is a member of a kinase family and is involved in the reorganization of the cytoskeleton. Mol. Cell Biol. 16: 5313-5327. 8816443
Ly, D., Resch, E., Ordiway, G. and DiNardo, S. (2017). Asymmetrically deployed actomyosin-based contractility generates a boundary between developing leg segments in Drosophila. Dev Biol 429(1): 165-176. PubMed ID: 28689737
Maekawa, M., et al. (1999). Signaling from Rho to the actin cytoskeleton through protein kinases ROCK and LIM-kinase. Science 285(5429): 895-8. PubMed Citation: 10436159
Marlow, G., et al. (2002). Zebrafish Rho kinase 2 acts downstream of Wnt11 to mediate cell polarity and effective convergence and extension movements. Curr. Biol. 12: 876-884. 12062050
McMullan, R., et al. (2004). Keratinocyte differentiation is regulated by the Rho and ROCK signaling pathway. Curr. Biol. 13: 2185-2189. 14680635
Mizuno, T., Amano, M., Kaibuchi, K. and Nishida, Y. (1999). Identification and characterization of Drosophila homolog of Rho-kinase. Gene 238(2): 437-444. 10570971
Mizuno, T., Tsutsui, K. and Nishida, Y. (2002). Drosophila myosin phosphatase and its role in dorsal closure. Development 129: 1215-1223. 11874917
Nakayama, M., et al. (2008). Rho-kinase phosphorylates PAR-3 and disrupts PAR complex formation. Dev. Cell 14(2): 205-15. PubMed Citation: 18267089
Neubueser, D. and Hipfner, D. R. (2010). Overlapping roles of Drosophila Drak and Rok kinases in epithelial tissue morphogenesis. Mol Biol Cell 21: 2869-2879. PubMed ID: 20573980
Nikolaidou, K. K. and Barrett, K. (2004). A Rho GTPase signaling pathway is used reiteratively in epithelial folding and potentially selects the outcome of Rho activation. Curr. Biol. 14: 1822-1826. 15498489
Olazabal, I. M., et al. (2002). Rho-kinase and Myosin-II control phagocytic cup formation during CR, but Not FcgammaR, phagocytosis. Curr. Biol. 12: 1413-1418. 12194823
Ongusaha, P. P., et al. (2006). RhoE is a pro-survival p53 target gene that inhibits ROCK I-mediated apoptosis in response to genotoxic stress. Curr. Biol. 16: 2466-2472. Medline abstract: 17174923
Oude Weernink, P. A., et al. (2000). Stimulation of phosphatidylinositol-4-phosphate 5-kinase by Rho-kinase. J. Biol. Chem. 275(14): 10168-74. 10744700
Padash Barmchi, M., Rogers, S. and Hacker, U. (2005). DRhoGEF2 regulates actin organization and contractility in the Drosophila blastoderm embryo. J. Cell Biol. 168: 575-585. 15699213
Piekny, A. J., Wissmann, A. and Mains, P. E. (2000). Embryonic morphogenesis in Caenorhabditis elegans integrates the activity of LET-502 Rho-binding kinase, MEL-11 myosin phosphatase, DAF-2 insulin receptor and FEM-2 PP2c phosphatase. Genetics 156(4): 1671-89. 11102366
Piekny, A. J., et al. (2003). The Caenorhabditis elegans nonmuscle myosin genes nmy-1 and nmy-2 function as redundant components of the let-502/Rho-binding kinase and mel-11/myosin phosphatase pathway during embryonic morphogenesis. Development 130: 5695-5704. 14522875
Raghavan, S., Vaezi, A. and Fuchs, E. (2003). A role for alphaß1 integrins in focal adhesion function and polarized cytoskeletal dynamics. Dev. Cell 5: 415-427. 12967561
Riveline, D., Zamir, E., Balaban, N. Q., Schwarz, U. S., Ishizaki, T., Narumiya, S., Kam, Z., Geiger, B. and Bershadsky, A. D. (2001). Focal contacts as mechanosensors: externally applied local mechanical force induces growth of focal contacts by an mDia1-dependent and ROCK-independent mechanism. J. Cell Biol. 153: 1175-1186. 11402062
Robertson, F., Pinal, N., Fichelson, P. and Pichaud, F. (2012). Atonal and EGFR signalling orchestrate rok- and Drak-dependent adherens junction remodelling during ommatidia morphogenesis. Development 139: 3432-3441. PubMed ID: 22874916
Roovers, K. and Assoian, R. K. (2003a). Effects of rho kinase and actin stress fibers on sustained extracellular signal-regulated kinase activity and activation of G(1) phase cyclin-dependent kinases. Mol Cell Biol. 23(12): 4283-94. 12773570
Roovers, K., et al. (2003b). Nuclear translocation of LIM kinase mediates Rho-Rho kinase regulation of Cyclin D1 expression. Dev. Cell 5: 273-284. 12919678
Sahai, E., et al. (1999). Transformation mediated by RhoA requires activity of ROCK kinases. Curr. Biol. 9(3): 136-45. PubMed Citation: 10021386
Schmidt, M., et al. (1999). A role for rho-kinase in rho-controlled phospholipase D stimulation by the m3 muscarinic acetylcholine receptor. J. Biol. Chem. 274(21): 14648-54. 10329658
Simoes Sde, M., Blankenship, J. T., Weitz, O., Farrell, D. L., Tamada, M., Fernandez-Gonzalez, R. and Zallen, J. A. (2010). Rho-kinase directs Bazooka/Par-3 planar polarity during Drosophila axis elongation. Dev Cell 19: 377-388. PubMed ID: 20833361
Tanaka, H., Yamashita, T., Asada, M., Mizutani, S., Yoshikawa, H. and Tohyama, M. (2002). Cytoplasmic p21Cip1/Waf1 regulates neurite remodeling by inhibiting Rho-kinase activity. J. Cell. Biol. 158: 321-329. 12119358
Lee, A., 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. PubMed ID: 15075368
Vasquez, C. G., Tworoger, M., Martin, A. C. (2014). Dynamic myosin phosphorylation regulates contractile pulses and tissue integrity during epithelial morphogenesis. J Cell Biol 206: 435-450. PubMed ID: 25092658
Verdier, V., et al. (2006a). Drosophila Rho-kinase (DRok) is required for tissue morphogenesis in diverse compartments of the egg chamber during oogenesis. Dev. Biol. 297(2): 417-32. Medline abstract: 16887114
Verdier, V., Guang-Chao-Chena and Settleman, J. (2006b). Rho-kinase regulates tissue morphogenesis via non-muscle myosin and LIM-kinase during Drosophila development. BMC Dev. Biol. 6: 38. Medline abstract: 16882341
Wang, Y. and Riechmann, V. (2007). The role of the actomyosin cytoskeleton in coordination of tissue growth during Drosophila oogenesis. Curr Biol 17(15): 1349-55. Medline abstract: 17656094
Watanabe, N., et al. (1999). Cooperation between mDia1 and ROCK in Rho-induced actin reorganization. Nat. Cell Biol. 1: 136-143. PubMed Citation: 10559899
Wei1, L., et al. (2001). Rho kinases play an obligatory role in vertebrate embryonic organogenesis. Development 128: 2953-2962. 11532918
Winter, C. G., et al. (2001). Drosophila Rho-associated kinase (Drok) links Frizzled-mediated planar cell polarity signaling to the actin cytoskeleton. Cell 105: 81-91. 11301004
Wissmann, A., et al. (1997). Caenorhabditis elegans LET-502 is related to Rho-binding kinases and human myotonic dystrophy kinase and interacts genetically with a homolog of the regulatory subunit of smooth muscle myosin phosphatase to affect cell shape. Genes Dev. 11: 409-422. 9042856
Yashiro, H., Loza, A. J., Skeath, J. B. and Longmore, G. D. (2014). Rho1 regulates adherens junction remodeling by promoting recycling endosome formation through activation of Myosin II. Mol Biol Cell 25(19):2956-69. PubMed ID: 25079692
Yasui, Y., et al. (1998). Roles of Rho-associated kinase in cytokinesis; mutations in Rho-associated kinase phosphorylation sites impair cytokinetic segregation of glial filaments. J. Cell Biol. 143(5): 1249-58. 9832553
Zhang, X.-F., et al. (2003). Rho-dependent contractile responses in the neuronal growth cone are independent of classical peripheral retrograde actin flow. Neuron 40: 931-944. 14659092
Zhang, Y., et al. (2009). Rock2 controls TGFbeta signaling and inhibits mesoderm induction in zebrafish embryos. J. Cell Sci. 122(Pt 13): 2197-207. PubMed Citation: 19509062
Zhao, Z. and Rivkees, S. A. (2004). Rho-associated kinases play a role in endocardial cell differentiation and migration. Dev. Biol. 275: 183-191. 15464581
date revised: 10 October 2014
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