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The Role of Rho Family GTPases in Development: Lessons from Drosophila melanogaster
Molecular Cell Biology Research Communications 1, 87–94 (1999) Article ID mcbr.1999.0119, available online at http://www.idealibrary.com on
MINIREVIEW The Role of Rho Family GTPases in Development: Lessons from Drosophila melanogaster Yu Lu and Jeffrey Settleman Massachusetts General Hospital Cancer Center and Harvard Medical School, Building 149, 13th Street, Charlestown, Massachusetts 02129
Received March 15, 1999
for a variety of cellular functions in most cell types, their precise mechanism of action and the organization of their associated signal transduction pathways are just beginning to be elucidated.
The Rho Subfamily of Small GTPases The Rho family of Ras-related GTPases comprises an expanding group of small GTP-binding proteins that regulate a variety of cellular functions (1). Genes encoding Rho-related proteins have been identified in several organisms, including mammals, insects, plants, and yeast, indicating that they have been highly conserved throughout evolution. Like their Ras counterparts, these GTPases function as molecular switches, cycling between active GTP-bound and inactive GDP-bound states to mediate intracellular signal transduction pathways in response to extracellular stimuli. Functional studies of the three prototype Rho family GTPases, Rho, Rac, and Cdc42, have revealed that these proteins perform essential functions in a variety of biological processes, including cell cycle progression and gene transcription. However, their most intensively studied function is that of cytoskeletal regulation (2). Several of the Rho family proteins can regulate actin cytoskeleton remodeling in response to extracellular signals. When microinjected into fibroblasts, activated Rho stimulates the formation of actin stress fibers and focal adhesions (3), Rac induces membrane ruffling (lamellipodia formation) (4), and Cdc42 promotes the formation of filopodia (5). Moreover, inhibiting particular Rho family GTPases in a variety of experimental systems blocks certain cellular response to extracellular stimuli, confirming their critical function as signal transduction intermediates. As potent regulators of the actin cytoskeleton, these proteins have also been linked to cellular processes that determine cell shape, motility, and adhesive properties (2). Thus, it has been suggested that the Rho GTPases are likely to be important regulators of the numerous morphogenetic events associated with the development of multicellular organisms. Although it is generally appreciated that Rho GTPases are critical switching components
Biochemical Regulation of Rho GTPases Much like their Ras counterparts, Rho GTPases function as molecular switches, and their ability to cycle between active and inactive states is tightly regulated (1). While their intrinsic GTPase activities are relatively slow, the hydrolysis of bound GTP by activated Rho proteins can be greatly accelerated by a family of proteins called GTPase activating proteins (GAPs) (1). Thus, the GAPs promote GTPase inactivation. Activation of the Rho GTPases is promoted by a class of proteins named guanine nucleotide exchange factors (GNEFs). The GNEFs appear to function by stabilizing a nucleotide-free form of the GTPase, such that it can bind GTP (which is considerably more abundant in the cell than GDP), thereby becoming activated (1). Together, these regulatory proteins provide stringent control over the nucleotide state of the GTPases, and appear to link the activation of the Rho GTPases to cell surface signals through mechanisms that are just beginning to be elucidated. Rho GTPase-Associated Signaling Pathways Upon activation to the GTP-bound form, GTPases undergo a conformational change that allows them to interact with so-called downstream effector targets, which contribute to the cellular response to GTPase activation. During the past few years, numerous proteins have been identified that bind specifically to the active GTP-bound Rho GTPases. Many of these proteins exhibit specific interactions with a particular Rho GTPase, although a few of them appear to be shared among different Rho proteins (1). Thus, it has been 87
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In general, this system, in which many of the components of mammalian signaling pathways appear to be well conserved, provides a setting in which loss-offunction and gain-of-function of particular genes in a variety of developmental processes can be directly examined. The ability both to easily generate transgenic fly lines and to perform genetic interaction studies, as well as mutational “modifier” screens, has provided investigators with powerful tools for dissecting signaling pathways in vivo. In the following sections, recent studies will be summarized in which the Drosophila system has been used to examine the regulation and function of the Rho family GTPases in a variety of developmental processes. As described below, these studies have revealed the power of this system in elucidating the role of the Rho GTPases and their associated signaling pathways in regulating the morphogenetic events required for the normal development of multicellular organisms.
difficult to establish the mechanisms by which signaling specificity is achieved in vivo. Many of the putative GTPase effector targets are protein kinases. For example, the Rho GTPase associates specifically with several identified protein kinases, including the PKC-related PKN (6, 7) and PRK2 kinases (8, 9), the ROK (Rho Kinase) family of serine/threonine kinases (10), and the Citron kinase (11). The closely related Rac and Cdc42 GTPases associate with a distinct family of kinases referred to as the PAK (p21-activated) kinases (12). These kinases are all moderately activated upon binding to the GTP bound forms of the respective GTPases, suggesting that upon GTPase activation, a signal is transduced to these protein kinases that leads to the phosphorylation of substrates. However, for most of these kinases, the relevant substrate targets remain unidentified. In addition to these protein kinases, several lipid kinases have also been identified as putative Rho effector targets (13–15). Much less is known about the upstream signaling pathways that lead to activation of the Rho GTPases. Rho proteins can be activated by various extracellular stimuli, such as LPA (lysophosphatidic acid), growth factors, bombesin and bradykinin (1). The LPA, bradykinin and bombesin receptors belong to the seventransmembrane-domain family and are linked to heterotrimeric G proteins, which may be required for activating Rho GTPases. The Rho/Rac GNEFs appear to be regulated by a variety of factors, including phosphorylation, lipid interactions, and membrane localization, indicating that diverse regulatory inputs may be utilized to promote their ability to activate the Rho family GTPases (1). Despite these observations, the signaling pathways leading from cell surface receptor activation to Rho/Rac/Cdc42 GTPase activation are largely unknown.
The Drosophila Rho GTPases and Their Expression in Embryogenesis At least five Drosophila Rho GTPases have been identified thus far (17–20). These include Rho1, Rac1, Rac2, Cdc42, and RhoL, which are 70-90% identical in amino acid sequence to their mammalian counterparts. These GTPases are expressed throughout embryogenesis, and some are widely expressed in many tissues (such as Rho1 and Rac2) while others exhibit more restricted expression in the mesoderm, gut, and nervous system later in development (such as Rac1 and Cdc42) (17, 18). This is consistent with the notion that Rho GTPases are important regulators of actin cytoskeletal changes required for the numerous morphogenetic events in Drosophila development. In the next sections, studies addressing the role of Rho family GTPases in several of the well studies aspects of Drosophila development are summarized.
Genetic Analysis of Rho GTPase Signaling in Drosophila
Oogenesis
As described above, biochemical studies in mammalian systems have yielded substantial information regarding the biochemical regulation of Rho GTPases, and the identification of many of their downstream targets. However, the biological significance of these findings, in most cases, remains somewhat elusive. As an alternative approach, genetic analyses of the evolutionarily conserved Rho GTPases in simpler organisms, such as Drosophila melanogaster, have begun to provide additional clues to their function. Drosophila, a model organism that is ideally suited to genetic and developmental studies, has been particularly useful in deciphering the organization of the Ras-mediated signal transduction pathway (16). Recently, this system has proved to be equally as powerful a tool to study Rho-associated signaling pathways in an in vivo context.
Drosophila oogenesis takes place in an organ structure referred to as the egg chamber. Each egg chamber consists of 15 nurse cells and a single oocyte that are connected through actin-rich structures called ring canals, and the 16-cell germ line is covered by a single layer of somatic cells known as follicle cells. In addition to the ring canals, the cortical regions of all germ cells and the adherent junction type connections between the germ cells and follicle cells also contain high concentrations of filamentous actin (21). During oogenesis, nurse cells eventually “dump” all of their cytosolic contents to the oocyte through the ring canals using an actin-myosin contractile system, while the follicle cells undergo characteristic shape changes and migrations that may be cytoskeleton-mediated. Therefore, it was predicted that Rho GTPases play a role in oogenesis. 88
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In one study, dominant-negative forms of Rac1, Cdc42 and RhoL were specifically expressed in a specialized subset of follicle cells (border cells) which normally migrate to the anterior tip of the egg chamber midway through oogenesis (19). While no requirement for Cdc42 or RhoL was observed, expression of dominant-negative Rac1 nearly completely blocked border cell migration without affecting other follicle cells, suggesting a specific role of Rac1 in this process. In the same study, it was demonstrated that while all three GTPases are required for transfer of nurse cell contents to oocytes, they each perform additional cell type specific functions during oogenesis. Gastrulation Drosophila gastrulation commences immediately following the cellularization of the syncitial blastoderm. This evolutionary conserved developmental process involves major morphogenetic events including ventral furrow formation and midgut invagination (22). Ventral furrow formation is largely dependent on cytoskeleton-mediated cell shape changes within a group of mesodermal precursors located along the length of the mid-ventral surface of the embryo, and results in the internalization of the presumptive mesoderm. In slightly later developmental stages, similar cell shape changes are responsible for the invaginations of anterior and posterior endodermal primordia, which subsequently give rise to the midgut. These cell shape changes involve actin cytoskeleton-mediated constriction of the apical membranes followed by a cell-shortening event (23). Recent studies revealed that a Rho GTPase signal transduction pathway controls the gastrulation process (24, 25). Specifically, a Drosophila RhoGEF (dRhoGEF2) was identified that was found to be essential for cell shape changes in gastrulation. Furthermore, expression of a dominant-negative Rho1 mutant in early embryos blocks ventral furrow formation as well as midgut invagination, suggesting that activation of Rho1 by dRhoGEF2 is essential for cell shape changes required for gastrulation (24). Two previously identified Drosophila gastrulation mutants, folded gastrulation (fog) (26) and concertina (cta) (27), exhibit very similar defects to that of dRhoGEF2. Interestingly, Fog is an extracellular ligand for an unidentified membrane receptor and Cta is a G subunit of a heterotrimeric G protein that may be coupled to the Fog receptor. It is tempting to speculate that Fog activates a receptor coupled G-protein containing Cta, which in turn activates the down stream Rho1 GTPase through dRhoGEF2 and ultimately leads to cell shape changes in gastrulation. Indeed, ectopic fog expression induces cell shape changes that can be blocked in either the cta (28) or dRhoGEF2 mutant background (24). An analogous pathway has been described in mammalian cells in which LPA activates the LPA receptor coupled
FIG. 1. Evolutionary conserved Rho-mediated signal transduction pathways for cell shape changes required for Drosophila gastrulation (left panel) and neuronal cell shape changes in mammals (right panel).
G-protein, leading to subsequent activation of the Rho GTPase via a RhoGEF (29), indicating that this pathway of Rho activation has been conserved evolutionarily (Fig. 1). Dorsal Closure Dorsal closure in Drosophila development occurs midway through embryogenesis. It is a major morphogenetic event in which embryonic epidermal cells stretch along their dorsal-ventral axis. Consequently, the two lateral epidermal cell sheets eventually meet at the dorsal midline, thereby closing the dorsal side of the embryo. The process involves neither cell division nor migration, and is solely dependent on cytoskeletonmediated cell shape changes within a subset of embryonic cells (30). Defects in dorsal closure result in a hole in the dorsal side of the larval cuticle, and can easily be detected. Recently, the Rho family proteins and their associated pathways have been implicated in dorsal closure (31–33). The mutant phenotype of one of the Rho GTPases has been described. Rho1 null allele mutants display a dorsal hole in the larval cuticle (33) (Fig. 2). Similar defects are seen in embryos expressing dominantnegative Rac1 (31) or Cdc42 (32) using a heat-shock promoter, suggesting a role for all three of these GTPases. Interestingly, mutants of several kinase pathway genes that function downstream of the respective Rho GTPases in mammalian system also exhibit dorsal closure defects. For example, mutations in the JNK cascade kinases, including hemipterous (the Jun N-terminal Kinase Kinase) (34) and basket (JNK) (Fig. 89
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tial muscle fibers (45). This fusion process requires assembly of actin filaments (46), suggesting that Rho GTPases might be involved. Indeed, one study using transgenic flies expressing mutant forms of Rho GTPases has revealed a role for Rac in myoblast fusion (18). Expression of dominant-negative Rac1 initially delays the fusion process, followed by excessive fusion of myoblasts, while constitutively active Rac1 completely blocks fusion, indicating that precise control of the Rac activity is essential for normal myoblast fusion. Despite a high degree of structural similarity between Rac and Cdc42, similar experiments using Cdc42 transgenic flies did not reveal a role of Cdc42 in myoblast fusion, indicating that this is a Rac specific function and that closely related Rho GTPases perform different biological functions. The signaling pathway involved in the Rac-mediated myoblast fusion is unknown. However, it was found that the Rac regulator, mbc, is also required for myoblast fusion (43). Although it is unclear from the Mbc protein sequence how Rac might be activated by Mbc, it is known that the mammalian Mbc homologue, DOCK180, associates with the adapter protein Crk (47), which binds the RacGEF, Vav (48). This raises the possibility that a multi-protein complex containing Mbc might function to recruit Rac to its GNEF, thereby activating the Rac mediated signaling pathway(s) required for both myoblast fusion and dorsal closure.
FIG. 2. Cuticle preparations of (A) wild type, (B) Rho1 72O (a null allele of Drosophila Rho1), and (C) bsk 1 (a strong loss-of-function allele of Drosophila JNK), showing dorsal closure defects in basket (bsk) and Rho1 mutant embryos.
2) (32, 35), their downstream transcription factors DJun (36 –39) and DFos (kayak) (40, 41), and the DJun target gene puckered (JNK phosphatase) (42), all result in dorsal closure defects. Recent studies have established a dorsal closure model in which Rac and/or Cdc42 signals through the JNK cascade to induce expression in the leading edge cells (the dorsal-most role of epidermal cells known to initiate the dorsal-ventral stretching of the two epidermal cell sheets) of Dpp, a secreted TGF-b-like ligand. It appears that secreted Dpp then signals the stretching of the more lateral epidermal cells, resulting in the movement of the two cell sheets toward the midline (36 –39). Thus far, the precise role of Rho1 in this process has not been determined. The identity of signaling components that initiate the dorsal closure process is still unclear. In particular, the pathways by which the Rho and Rac GTPases are activated to initiate the stretching of epidermal cells, which is required for dorsal closure, have not yet been elucidated. However, it is likely that GNEFs will be involved, as well as membrane receptors and/or adapters. One such candidate is mbc (myoblast city), a gene which is also required for dorsal closure and appears to function as a Rac-specific upstream regulator of the dorsal closure process (43, 44). Mbc was identified in a mutational modifier screen designed to identify specific components of the Rac signaling pathway, and the human Mbc homologue, DOCK180, was subsequently found to function as an upstream regulator of Rac activity (44). Thus, the genetic approach was able to reveal the function of a human protein whose cellular role was previously unknown.
Neural Development The neuronal growth cone is a highly dynamic actinrich structure located at the tip of a growing axon. The organized rearrangement of axonal actin structures is reminiscent of the Rac induced lamellipodia and Cdc42 induced filopodia seen in fibroblasts; moreover, high expression levels of both Rac and Cdc42 in the Drosophila nervous system have been observed during late stages of embryogenesis (18). Therefore, a role for Rac or Cdc42 in axon outgrowth and guidance during development is expected. Expression of dominantnegative Rac1 in neuroblasts or in the mature central and peripheral nervous system caused embryonic lethality associated with severe loss of axons (but not dendrites), suggesting a specific role of Rac in axon outgrowth initiation (18). Similar but more severe axonal losses were observed when activated Rac was expressed. In addition, by expressing activated Rac at slightly later stages of neuronal development so that axon outgrowth is initiated, it was determined that proper Rac activation is required for axon elongation. Expression of mutant forms of Cdc42 was found to cause qualitatively different neuronal defects (18). Notably, activated Cdc42 inhibited both axon and dendrite outgrowth. Moreover, affects on neuronal position in addition to axon outgrowth were observed, suggest-
Muscle Development As in vertebrates, Drosophila muscle development involves the fusion of myoblasts to form mature synci90
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FIG. 3. Scanning electron microscopy images of Drosophila eyes from (A) wild type, (B) pGMR-Rho1, (C) pGMR-Rac1, and (D) pGMR-Cdc42 transgenic flies, showing the “rough-eye” phenotypes in transgenic flies that overexpress Rho1, Rac1, and Cdc42 in Drosophila eyes, respectively.
ing an additional role of Cdc42 in neuronal migration. Further studies are required to examine the defects associated with the loss-of-function mutants of both GTPases before firm conclusions regarding the normal roles of Rac and Cdc42 in neuronal development can be drawn. The organization of Rac and Cdc42 mediated signal transduction pathways in Drosophila neuronal development remain largely unknown. However, a few studies suggest that some of the same types of components used in other Rho signaling pathways are involved. For example, a Drosophila RhoGEF, still life, was recently identified in a genetic screen searching for motor activity-defective mutants (49). Still life expression in late embryogenesis is restricted in neurons, predomi-
nantly in the synaptic regions, suggesting a role in synaptic function, and still life mutant flies exhibit reduced locomotion. Transgenic flies expressing a putative activated form of the Still life protein in neurons display defects in motor axon elongation and formation of synaptic arbors on target muscles. In another report, expression of a dominant-negative form of Rac1 in neurons caused axons associated with the intersegmental nerve b to bypass and extend beyond their normal synaptic targets (50). This “bypass” phenotype is enhanced by the absence of the receptor-tyrosine phosphatase, DLAR, which has also been implicated in motor axon guidance (51). Since the mammalian homologue of DLAR interacts directly with Trio, a protein that contains two Rac/Rho GEF-like domains (52), it is 91
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Excessive activity of the Rho GTPases perturbs the normal development of the eye (17). When overexpressed in transgenic lines, wild type forms of Rho1, Rac1, Rac2, and Cdc42 can each disrupt the normal ommatidial structure of the eye and result in an externally “rough” phenotype (Fig. 3). For example, expression of a Rho1 transgene causes a rough-eye phenotype associated with disruption of the morphology of the photoreceptors (17). In addition, the lattice formed by secondary and tertiary pigment cells as well as the general ommatidial architecture is completely disrupted. Furthermore, disruption of the normal appearance of the polymerized actin in pupal eye discs from these transgenics suggests that the ability of Rho1 to affect actin organization plays an important role in eye morphogenesis. Expression of Rac1, Rac2, and Cdc42 GTPases in the eye similarly disrupts normal eye development, although the observed defects are different in each case, indicating that this system is useful for distinguishing the activities of closely related Rho GTPases in vivo (44).
embryonic ectoderm, and become the imaginal discs during larval development. These discs eventually give rise to all adult tissues. Aside from the commonly observed apical-basal polarity in epithelial tissue culture cells, epithelial cells can also develop a planar polarity relative to the body axis (53). This is reflected in the appearance of distally pointed hairs on the Drosophila wing blades and the regular arrays of trapezoidal shaped rhabdomeres in the Drosophila compound eye. To study the role of Rho GTPases in establishing tissue polarity in the fly wing disc, mutant forms of Rac1 and Cdc42 were expressed specifically in wing discs (54). From those experiments, it was found that Rac1 activity was essential for the proper assembly of cell adherens junctions as well as for the establishment of planar polarity, while Cdc42 was found to be required for epithelial cell shape changes, but is not required for actin assembly at adherens junctions. Neither of these GTPases was required to maintain polarity once it was established. In addition, detailed examination revealed distinct functions for Rac and Cdc42 in regulating different aspects of wing hair formation (55). Specifically, expression of dominant-negative Cdc42 stunts wing hair growth, leading to a failure of wing hair cells to accumulate actin in their distal regions. This suggests that Cdc42 might be responsible for polarized membrane outgrowth which is analogous to the Cdc42 stimulated filopodia formation in fibroblasts. On the other hand, dominant-negative Rac1 promotes the outgrowth of multiple hairs from a single cell, which also exhibit gaps in junctional actin and a disorganized apical microtubule web, indicating that Rac1 is responsible for both actin polymerization and microtubule organization. The existing Rho1 mutant further confirmed the role of Rho GTPases in establishing tissue polarity in wing development (33). Wing clones containing hypomorphic Rho1 alleles exhibit abnormal wing hair polarity. Similarly, in Rho1 eye clones, a large number of ommatidia were incorrectly oriented while the relative position of photoreceptors and accessory cells remain unaltered. This is the same phenotype seen in two previously identified tissue polarity mutants: frizzled and disheveled. Frizzled appears to be a G-protein coupled receptor, and disheveled is a cytoplasmic signaling molecule. Genetic experiments established that Rho1 functions downstream of the frizzled-disheveled mediated tissue polarity pathway (33), which is consistent with the general theme linking G-protein coupled receptors to the Rho signaling pathway.
Tissue Polarity
Summary
Polarized epithelial cells perform a variety of specialized functions and epithelial polarity plays an important role in Drosophila morphogenesis. Drosophila epithelial cells are derived from the invaginated
It has become increasingly clear in the last few years that the Rho family GTPases regulate cytoskeleton rearrangements that are essential for a variety of morphogenetic events associated with the development of
TABLE I
Summary of Rho GTPase Signaling Components Involved in Various Drosophila Developmental Processes Developmental processes Oogenesis: border cell migration Oogenesis: transfer of nurse cell contents to oocytes Gastrulation Dorsal closure
Muscle development Neural development Eye development Tissue polarity: wing development Tissue polarity: eye development
Genes/Pathways involved
References
Rac1
(19)
Rac1, Cdc42, RhoL
(19)
Fog, cta, DRhoGEF2, Rho1 Rho1, Rac1, Cdc42, JNK pathway genes, Dpp pathway genes, and mbc Rac1, mbc Rac1, Cdc42, still life Rho1, Rac1, Rac2, Cdc42 Rac1, Cdc42, Rho1
(24–27)
Rho1, frizzled, disheveled
(31–44)
(18, 43) (18, 49, 50) (17, 44) (33, 54, 55) (33)
possible that DLAR activates Rac1 through a Trio-like GEF to mediate an axon guidance signal. Eye Development
92
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MOLECULAR CELL BIOLOGY RESEARCH COMMUNICATIONS 18b.Luo, L., Liao, Y. J., Jan, L. Y., and Jan, Y. N. (1994) Genes Dev. 8, 1787–1802. 19. Murphy, A. M., and Montell, D. J. (1996) J. Cell Biol. 133, 617– 630. 20. Sasamura, T., Kobayashi, T., Kojima, S., Qadota, H., Ohya, Y., Masai, I., and Hotta, Y. (1997) Mol. Gen. Genet. 254, 486 – 494. 21. Cooley, L., and Theurkauf, W. E. (1994) Science 266, 590 –596. 22. Sweeton, D., Parks, S., Costa, M., and Wieschaus, E. (1991) Development 112, 775–789. 23. Leptin, M. (1995) Annu. Rev. Cell Dev. Biol. 11, 189 –212. 24. Barrett, K., Leptin, M., and Settleman, J. (1997) Cell 91, 905– 915. 25. Hacker, U., and Perrimon, N. (1998) Genes Dev. 12, 274 –284. 26. Costa, M., Wilson, E. T., and Wieschaus, E. (1994) Cell 76, 1075–1089. 27. Parks, S., and Wieschaus, E. (1991) Cell 64, 447– 458. 28. Morize, P., Christiansen, A., Costa, M., Parks, S., and Wieschaus, E. (1998) Dev. Suppl. 125, 589 –597. 29. Gebbink, M. F., Kranenburg, O., Poland, M., van Horck, F. P., Houssa, B., and Moolenaar, W. H. (1997) J. Cell. Biol. 137, 1603–1613. 30. Martinez-Arias, A. (1993) in The Development of Drosophila melanogaster (Bate, M., and Martinez-Arias, A., Eds.), pp. 517– 608. Cold Spring Harbor Laboratory Press, New York, NY. 31. Harden, N., Loh, H. Y., Chia, W., and Lim, L. (1995) Development 121, 903–914. 32. Riesgo-Escovar, J. R., Jenni, M., Fritz, A., and Hafen, E. (1996) Genes Dev. 10, 2759 –2768. 33. Strutt, D. I., Weber, U., and Mlodzik, M. (1997) Nature 387, 292–295. 34. Glise, B., Bourbon, H., and Noselli, S. (1995) Cell 83, 451– 461. 35. Sluss, H. K., Han, Z., Barrett, T., Davis, R. J., and Ip, Y. T. (1996) Genes Dev. 10, 2745–2758. 36. Glise, B., and Noselli, S. (1997) Genes Dev. 11, 1738 –1747. 37. Hou, X. S., Goldstein, E. S., and Perrimon, N. (1997) Genes Dev. 11, 1728 –1737. 38. Kockel, L., Zeitlinger, J., Staszewski, L. M., Mlodzik, M., and Bohmann, D. (1997) Genes Dev. 11, 1748 –1758. 39. Riesgo-Escovar, J. R., and Hafen, E. (1997) Genes Dev. 11, 1717– 1727. 40. Riesgo-Escovar, J. R., and Hafen, E. (1997) Science 278, 669 – 672. 41. Zeitlinger, J., Kockel, L., Peverali, F. A., Jackson, D. B., Mlodzik, M., and Bohmann, D. (1997) EMBO J. 16, 7393–7401. 42. Martin-Blanco, E., Gampel, A., Ring, J., Virdee, K., Kirov, N., Tolkovsky, A. M., and Martinez-Arias, A. (1998) Genes Dev. 12, 557–570. 43. Erickson, M. R., Galletta, B. J., and Abmayr, S. M. (1997) J. Cell Biol. 138, 589 – 603. 44. Nolan, K. M., Barrett, K., Lu, Y., Hu, K. Q., Vincent, S., and Settleman, J. (1998) Genes Dev. 12, 3337–3342. 45. Bate, M. (1993) in The Development of Drosophila melanogaster (Bate, M., and Martinez-Arias, A., Eds.), pp. 1013–1090. Cold Spring Harbor Laboratory Press, New York, NY. 46. Holtzer, H., Strahs, K., Biehl, J., Somlyo, A. P., and Ishikawa, H. (1975) Science 188, 943–945. 47. Hasegawa, H., Kiyokawa, E., Tanaka, S., Nagashima, K., Gotoh, N., Shibuya, M., Kurata, T., and Matsuda, M. (1996) Mol. Cell. Biol. 16, 1770 –1776. 48. Smit, L., van der Horst, G., and Borst, J. (1996) J. Biol. Chem. 271, 8564 – 8569. 49. Sone, M., Hoshino, M., Suzuki, E., Kuroda, S., Kaibuchi, K.,
multicellular organisms. In particular, Drosophila has provided an excellent in vivo system for deciphering the signaling pathways mediated by Rho GTPases, as well as establishing the role of these pathways in numerous developmental processes (Table I). Continued use of this system will undoubtedly lead to the identification of additional Rho signaling components and information regarding the function and organization of the Rho signaling pathways in tissue morphogenesis. The striking similarity between Drosophila and mammalian Rho signaling components identified thus far indicates that the Rho pathways are highly conserved in evolution. Therefore, the findings from the Drosophila system can be extrapolated to higher organisms, including humans. Combined with the rapid progress in the human and Drosophila genome projects, these findings should contribute greatly to our understanding of mammalian Rho GTPase signaling pathways and their roles in normal development and pathological conditions. REFERENCES 1. Van Aelst, L., and D’Souza-Schorey, C. (1997) Genes Dev. 11, 2295–2322. 2. Hall, A. (1998) Science 280, 2074 –2075. 3. Ridley, A. J., and Hall, A. (1992) Cell 70, 389 –399. 4. Ridley, A. J., Paterson, H. F., Johnston, C. L., Diekmann, D., and Hall, A. (1992) Cell 70, 401– 410. 5. Nobes, C. D., and Hall, A. (1995) Cell 81, 53– 62. 6. Watanabe, G., Saito, Y., Madaule, P., Ischizaki, T., Fujisawa, K., Morii, N., Mukai, H., Ono, Y., Kakizuka, S., and Narumiya, S. (1996) Science 271, 645– 648. 7. Amano, M., Mukai, H., Ono, Y., Chihara, K., Matsui, T., Hamajima, Y., Okawa, K., Iwamatsu, A., and Kaibuchi, K. (1996) Science 271, 648 – 650. 8. Quilliam, L. A., Lambert, Q. T., Mickelson-Young, L. A., Westwick, J. K., Sparks, A. B., Kay, B. K., Jenkins, N. A., 9. Gilbert, D. J., Copeland, N. G., and Der, C. J. (1996) J. Biol. Chem. 271, 28772–28776. 10. Vincent, S., and Settleman, J. (1997) Mol. Cell. Biol. 17, 2247– 2256. 11. Leung, T., Chen, X.-Q., Manser, E., and Lim, L. (1996) Mol. Cell. Biol. 16, 5313–5327. 12. Madaule, P., Eda, M., Watanabe, N., Fujisawa, K., Matsuoka, T., Bito, H., Ishizaki, T., and Narumiya, S. (1998) Nature 394, 491– 494. 13. Bagrodia, S., Taylor, S. J., Creasy, C. L., Chernoff, J., and Cerione, R. A. (1995) J. Biol. Chem. 270, 22731–22737. 14. Chong, L. D., Traynor-Kaplan, A., Bokoch, G. M., and Schwartz, M. A. (1994) Cell 79, 507–513. 15. Zhang, J., King, W. G., Dillon, S., Hall, A., Feig, L., and Rittenhouse, S. E. (1993) J. Biol. Chem. 268, 22251–22254. 16. Zheng, Y., Bagrodia, S., and Cerione, R. A. (1994) J. Biol. Chem. 269, 18727–18730. 17. Simon, M. A., Bowtell, D. D. L., Dodson, G. S., Laverty, T. R., and Rubin, G. M. (1991) Cell 67, 701–716. 18a.Hariharan, I. K., Hu, K. Q., Asha, H., Quintanilla, A., Ezzell, R. M., and Settleman, J. (1995) EMBO Journal 14, 292–302. 93
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Nakagoshi, H., Saigo, K., Nabeshima, Y., and Hama, C. (1997) Science 275, 543–547. 50. Kaufmann, N., Wills, Z. P., and Van Vactor, D. (1998) Development 125, 453– 461. 51. Krueger, N. X., Van Vactor, D., Wan, H. I., Gelbart, W. M., Goodman, C. S., and Saito, H. (1996) Cell 84, 611– 622. 52. Debant, A., Serra-Pages, C., Seipel, K., O’Brien, S., Tang, M.,
Park, S. H., and Streuli, M. (1996) Proc. Natl. Acad. Sci. USA 93, 5466 –5471. 53. Eaton, S. (1997) Curr. Opin. Cell Biol. 9, 860 – 866. 54. Eaton, S., Auvinen, P., Luo, L., Jan, Y. N., and Simons, K. (1995) J. Cell Biol. 131, 151–164. 55. Eaton, S., Wepf, R., and Simons, K. (1996) J. Cell Biol. 135, 1277–1289.
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MINIREVIEW The Role of Rho Family GTPases in Development: Lessons from Drosophila melanogaster Yu Lu and Jeffrey Settleman Massachusetts General Hospital Cancer Center and Harvard Medical School, Building 149, 13th Street, Charlestown, Massachusetts 02129
Received March 15, 1999
for a variety of cellular functions in most cell types, their precise mechanism of action and the organization of their associated signal transduction pathways are just beginning to be elucidated.
The Rho Subfamily of Small GTPases The Rho family of Ras-related GTPases comprises an expanding group of small GTP-binding proteins that regulate a variety of cellular functions (1). Genes encoding Rho-related proteins have been identified in several organisms, including mammals, insects, plants, and yeast, indicating that they have been highly conserved throughout evolution. Like their Ras counterparts, these GTPases function as molecular switches, cycling between active GTP-bound and inactive GDP-bound states to mediate intracellular signal transduction pathways in response to extracellular stimuli. Functional studies of the three prototype Rho family GTPases, Rho, Rac, and Cdc42, have revealed that these proteins perform essential functions in a variety of biological processes, including cell cycle progression and gene transcription. However, their most intensively studied function is that of cytoskeletal regulation (2). Several of the Rho family proteins can regulate actin cytoskeleton remodeling in response to extracellular signals. When microinjected into fibroblasts, activated Rho stimulates the formation of actin stress fibers and focal adhesions (3), Rac induces membrane ruffling (lamellipodia formation) (4), and Cdc42 promotes the formation of filopodia (5). Moreover, inhibiting particular Rho family GTPases in a variety of experimental systems blocks certain cellular response to extracellular stimuli, confirming their critical function as signal transduction intermediates. As potent regulators of the actin cytoskeleton, these proteins have also been linked to cellular processes that determine cell shape, motility, and adhesive properties (2). Thus, it has been suggested that the Rho GTPases are likely to be important regulators of the numerous morphogenetic events associated with the development of multicellular organisms. Although it is generally appreciated that Rho GTPases are critical switching components
Biochemical Regulation of Rho GTPases Much like their Ras counterparts, Rho GTPases function as molecular switches, and their ability to cycle between active and inactive states is tightly regulated (1). While their intrinsic GTPase activities are relatively slow, the hydrolysis of bound GTP by activated Rho proteins can be greatly accelerated by a family of proteins called GTPase activating proteins (GAPs) (1). Thus, the GAPs promote GTPase inactivation. Activation of the Rho GTPases is promoted by a class of proteins named guanine nucleotide exchange factors (GNEFs). The GNEFs appear to function by stabilizing a nucleotide-free form of the GTPase, such that it can bind GTP (which is considerably more abundant in the cell than GDP), thereby becoming activated (1). Together, these regulatory proteins provide stringent control over the nucleotide state of the GTPases, and appear to link the activation of the Rho GTPases to cell surface signals through mechanisms that are just beginning to be elucidated. Rho GTPase-Associated Signaling Pathways Upon activation to the GTP-bound form, GTPases undergo a conformational change that allows them to interact with so-called downstream effector targets, which contribute to the cellular response to GTPase activation. During the past few years, numerous proteins have been identified that bind specifically to the active GTP-bound Rho GTPases. Many of these proteins exhibit specific interactions with a particular Rho GTPase, although a few of them appear to be shared among different Rho proteins (1). Thus, it has been 87
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In general, this system, in which many of the components of mammalian signaling pathways appear to be well conserved, provides a setting in which loss-offunction and gain-of-function of particular genes in a variety of developmental processes can be directly examined. The ability both to easily generate transgenic fly lines and to perform genetic interaction studies, as well as mutational “modifier” screens, has provided investigators with powerful tools for dissecting signaling pathways in vivo. In the following sections, recent studies will be summarized in which the Drosophila system has been used to examine the regulation and function of the Rho family GTPases in a variety of developmental processes. As described below, these studies have revealed the power of this system in elucidating the role of the Rho GTPases and their associated signaling pathways in regulating the morphogenetic events required for the normal development of multicellular organisms.
difficult to establish the mechanisms by which signaling specificity is achieved in vivo. Many of the putative GTPase effector targets are protein kinases. For example, the Rho GTPase associates specifically with several identified protein kinases, including the PKC-related PKN (6, 7) and PRK2 kinases (8, 9), the ROK (Rho Kinase) family of serine/threonine kinases (10), and the Citron kinase (11). The closely related Rac and Cdc42 GTPases associate with a distinct family of kinases referred to as the PAK (p21-activated) kinases (12). These kinases are all moderately activated upon binding to the GTP bound forms of the respective GTPases, suggesting that upon GTPase activation, a signal is transduced to these protein kinases that leads to the phosphorylation of substrates. However, for most of these kinases, the relevant substrate targets remain unidentified. In addition to these protein kinases, several lipid kinases have also been identified as putative Rho effector targets (13–15). Much less is known about the upstream signaling pathways that lead to activation of the Rho GTPases. Rho proteins can be activated by various extracellular stimuli, such as LPA (lysophosphatidic acid), growth factors, bombesin and bradykinin (1). The LPA, bradykinin and bombesin receptors belong to the seventransmembrane-domain family and are linked to heterotrimeric G proteins, which may be required for activating Rho GTPases. The Rho/Rac GNEFs appear to be regulated by a variety of factors, including phosphorylation, lipid interactions, and membrane localization, indicating that diverse regulatory inputs may be utilized to promote their ability to activate the Rho family GTPases (1). Despite these observations, the signaling pathways leading from cell surface receptor activation to Rho/Rac/Cdc42 GTPase activation are largely unknown.
The Drosophila Rho GTPases and Their Expression in Embryogenesis At least five Drosophila Rho GTPases have been identified thus far (17–20). These include Rho1, Rac1, Rac2, Cdc42, and RhoL, which are 70-90% identical in amino acid sequence to their mammalian counterparts. These GTPases are expressed throughout embryogenesis, and some are widely expressed in many tissues (such as Rho1 and Rac2) while others exhibit more restricted expression in the mesoderm, gut, and nervous system later in development (such as Rac1 and Cdc42) (17, 18). This is consistent with the notion that Rho GTPases are important regulators of actin cytoskeletal changes required for the numerous morphogenetic events in Drosophila development. In the next sections, studies addressing the role of Rho family GTPases in several of the well studies aspects of Drosophila development are summarized.
Genetic Analysis of Rho GTPase Signaling in Drosophila
Oogenesis
As described above, biochemical studies in mammalian systems have yielded substantial information regarding the biochemical regulation of Rho GTPases, and the identification of many of their downstream targets. However, the biological significance of these findings, in most cases, remains somewhat elusive. As an alternative approach, genetic analyses of the evolutionarily conserved Rho GTPases in simpler organisms, such as Drosophila melanogaster, have begun to provide additional clues to their function. Drosophila, a model organism that is ideally suited to genetic and developmental studies, has been particularly useful in deciphering the organization of the Ras-mediated signal transduction pathway (16). Recently, this system has proved to be equally as powerful a tool to study Rho-associated signaling pathways in an in vivo context.
Drosophila oogenesis takes place in an organ structure referred to as the egg chamber. Each egg chamber consists of 15 nurse cells and a single oocyte that are connected through actin-rich structures called ring canals, and the 16-cell germ line is covered by a single layer of somatic cells known as follicle cells. In addition to the ring canals, the cortical regions of all germ cells and the adherent junction type connections between the germ cells and follicle cells also contain high concentrations of filamentous actin (21). During oogenesis, nurse cells eventually “dump” all of their cytosolic contents to the oocyte through the ring canals using an actin-myosin contractile system, while the follicle cells undergo characteristic shape changes and migrations that may be cytoskeleton-mediated. Therefore, it was predicted that Rho GTPases play a role in oogenesis. 88
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In one study, dominant-negative forms of Rac1, Cdc42 and RhoL were specifically expressed in a specialized subset of follicle cells (border cells) which normally migrate to the anterior tip of the egg chamber midway through oogenesis (19). While no requirement for Cdc42 or RhoL was observed, expression of dominant-negative Rac1 nearly completely blocked border cell migration without affecting other follicle cells, suggesting a specific role of Rac1 in this process. In the same study, it was demonstrated that while all three GTPases are required for transfer of nurse cell contents to oocytes, they each perform additional cell type specific functions during oogenesis. Gastrulation Drosophila gastrulation commences immediately following the cellularization of the syncitial blastoderm. This evolutionary conserved developmental process involves major morphogenetic events including ventral furrow formation and midgut invagination (22). Ventral furrow formation is largely dependent on cytoskeleton-mediated cell shape changes within a group of mesodermal precursors located along the length of the mid-ventral surface of the embryo, and results in the internalization of the presumptive mesoderm. In slightly later developmental stages, similar cell shape changes are responsible for the invaginations of anterior and posterior endodermal primordia, which subsequently give rise to the midgut. These cell shape changes involve actin cytoskeleton-mediated constriction of the apical membranes followed by a cell-shortening event (23). Recent studies revealed that a Rho GTPase signal transduction pathway controls the gastrulation process (24, 25). Specifically, a Drosophila RhoGEF (dRhoGEF2) was identified that was found to be essential for cell shape changes in gastrulation. Furthermore, expression of a dominant-negative Rho1 mutant in early embryos blocks ventral furrow formation as well as midgut invagination, suggesting that activation of Rho1 by dRhoGEF2 is essential for cell shape changes required for gastrulation (24). Two previously identified Drosophila gastrulation mutants, folded gastrulation (fog) (26) and concertina (cta) (27), exhibit very similar defects to that of dRhoGEF2. Interestingly, Fog is an extracellular ligand for an unidentified membrane receptor and Cta is a G subunit of a heterotrimeric G protein that may be coupled to the Fog receptor. It is tempting to speculate that Fog activates a receptor coupled G-protein containing Cta, which in turn activates the down stream Rho1 GTPase through dRhoGEF2 and ultimately leads to cell shape changes in gastrulation. Indeed, ectopic fog expression induces cell shape changes that can be blocked in either the cta (28) or dRhoGEF2 mutant background (24). An analogous pathway has been described in mammalian cells in which LPA activates the LPA receptor coupled
FIG. 1. Evolutionary conserved Rho-mediated signal transduction pathways for cell shape changes required for Drosophila gastrulation (left panel) and neuronal cell shape changes in mammals (right panel).
G-protein, leading to subsequent activation of the Rho GTPase via a RhoGEF (29), indicating that this pathway of Rho activation has been conserved evolutionarily (Fig. 1). Dorsal Closure Dorsal closure in Drosophila development occurs midway through embryogenesis. It is a major morphogenetic event in which embryonic epidermal cells stretch along their dorsal-ventral axis. Consequently, the two lateral epidermal cell sheets eventually meet at the dorsal midline, thereby closing the dorsal side of the embryo. The process involves neither cell division nor migration, and is solely dependent on cytoskeletonmediated cell shape changes within a subset of embryonic cells (30). Defects in dorsal closure result in a hole in the dorsal side of the larval cuticle, and can easily be detected. Recently, the Rho family proteins and their associated pathways have been implicated in dorsal closure (31–33). The mutant phenotype of one of the Rho GTPases has been described. Rho1 null allele mutants display a dorsal hole in the larval cuticle (33) (Fig. 2). Similar defects are seen in embryos expressing dominantnegative Rac1 (31) or Cdc42 (32) using a heat-shock promoter, suggesting a role for all three of these GTPases. Interestingly, mutants of several kinase pathway genes that function downstream of the respective Rho GTPases in mammalian system also exhibit dorsal closure defects. For example, mutations in the JNK cascade kinases, including hemipterous (the Jun N-terminal Kinase Kinase) (34) and basket (JNK) (Fig. 89
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tial muscle fibers (45). This fusion process requires assembly of actin filaments (46), suggesting that Rho GTPases might be involved. Indeed, one study using transgenic flies expressing mutant forms of Rho GTPases has revealed a role for Rac in myoblast fusion (18). Expression of dominant-negative Rac1 initially delays the fusion process, followed by excessive fusion of myoblasts, while constitutively active Rac1 completely blocks fusion, indicating that precise control of the Rac activity is essential for normal myoblast fusion. Despite a high degree of structural similarity between Rac and Cdc42, similar experiments using Cdc42 transgenic flies did not reveal a role of Cdc42 in myoblast fusion, indicating that this is a Rac specific function and that closely related Rho GTPases perform different biological functions. The signaling pathway involved in the Rac-mediated myoblast fusion is unknown. However, it was found that the Rac regulator, mbc, is also required for myoblast fusion (43). Although it is unclear from the Mbc protein sequence how Rac might be activated by Mbc, it is known that the mammalian Mbc homologue, DOCK180, associates with the adapter protein Crk (47), which binds the RacGEF, Vav (48). This raises the possibility that a multi-protein complex containing Mbc might function to recruit Rac to its GNEF, thereby activating the Rac mediated signaling pathway(s) required for both myoblast fusion and dorsal closure.
FIG. 2. Cuticle preparations of (A) wild type, (B) Rho1 72O (a null allele of Drosophila Rho1), and (C) bsk 1 (a strong loss-of-function allele of Drosophila JNK), showing dorsal closure defects in basket (bsk) and Rho1 mutant embryos.
2) (32, 35), their downstream transcription factors DJun (36 –39) and DFos (kayak) (40, 41), and the DJun target gene puckered (JNK phosphatase) (42), all result in dorsal closure defects. Recent studies have established a dorsal closure model in which Rac and/or Cdc42 signals through the JNK cascade to induce expression in the leading edge cells (the dorsal-most role of epidermal cells known to initiate the dorsal-ventral stretching of the two epidermal cell sheets) of Dpp, a secreted TGF-b-like ligand. It appears that secreted Dpp then signals the stretching of the more lateral epidermal cells, resulting in the movement of the two cell sheets toward the midline (36 –39). Thus far, the precise role of Rho1 in this process has not been determined. The identity of signaling components that initiate the dorsal closure process is still unclear. In particular, the pathways by which the Rho and Rac GTPases are activated to initiate the stretching of epidermal cells, which is required for dorsal closure, have not yet been elucidated. However, it is likely that GNEFs will be involved, as well as membrane receptors and/or adapters. One such candidate is mbc (myoblast city), a gene which is also required for dorsal closure and appears to function as a Rac-specific upstream regulator of the dorsal closure process (43, 44). Mbc was identified in a mutational modifier screen designed to identify specific components of the Rac signaling pathway, and the human Mbc homologue, DOCK180, was subsequently found to function as an upstream regulator of Rac activity (44). Thus, the genetic approach was able to reveal the function of a human protein whose cellular role was previously unknown.
Neural Development The neuronal growth cone is a highly dynamic actinrich structure located at the tip of a growing axon. The organized rearrangement of axonal actin structures is reminiscent of the Rac induced lamellipodia and Cdc42 induced filopodia seen in fibroblasts; moreover, high expression levels of both Rac and Cdc42 in the Drosophila nervous system have been observed during late stages of embryogenesis (18). Therefore, a role for Rac or Cdc42 in axon outgrowth and guidance during development is expected. Expression of dominantnegative Rac1 in neuroblasts or in the mature central and peripheral nervous system caused embryonic lethality associated with severe loss of axons (but not dendrites), suggesting a specific role of Rac in axon outgrowth initiation (18). Similar but more severe axonal losses were observed when activated Rac was expressed. In addition, by expressing activated Rac at slightly later stages of neuronal development so that axon outgrowth is initiated, it was determined that proper Rac activation is required for axon elongation. Expression of mutant forms of Cdc42 was found to cause qualitatively different neuronal defects (18). Notably, activated Cdc42 inhibited both axon and dendrite outgrowth. Moreover, affects on neuronal position in addition to axon outgrowth were observed, suggest-
Muscle Development As in vertebrates, Drosophila muscle development involves the fusion of myoblasts to form mature synci90
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FIG. 3. Scanning electron microscopy images of Drosophila eyes from (A) wild type, (B) pGMR-Rho1, (C) pGMR-Rac1, and (D) pGMR-Cdc42 transgenic flies, showing the “rough-eye” phenotypes in transgenic flies that overexpress Rho1, Rac1, and Cdc42 in Drosophila eyes, respectively.
ing an additional role of Cdc42 in neuronal migration. Further studies are required to examine the defects associated with the loss-of-function mutants of both GTPases before firm conclusions regarding the normal roles of Rac and Cdc42 in neuronal development can be drawn. The organization of Rac and Cdc42 mediated signal transduction pathways in Drosophila neuronal development remain largely unknown. However, a few studies suggest that some of the same types of components used in other Rho signaling pathways are involved. For example, a Drosophila RhoGEF, still life, was recently identified in a genetic screen searching for motor activity-defective mutants (49). Still life expression in late embryogenesis is restricted in neurons, predomi-
nantly in the synaptic regions, suggesting a role in synaptic function, and still life mutant flies exhibit reduced locomotion. Transgenic flies expressing a putative activated form of the Still life protein in neurons display defects in motor axon elongation and formation of synaptic arbors on target muscles. In another report, expression of a dominant-negative form of Rac1 in neurons caused axons associated with the intersegmental nerve b to bypass and extend beyond their normal synaptic targets (50). This “bypass” phenotype is enhanced by the absence of the receptor-tyrosine phosphatase, DLAR, which has also been implicated in motor axon guidance (51). Since the mammalian homologue of DLAR interacts directly with Trio, a protein that contains two Rac/Rho GEF-like domains (52), it is 91
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Excessive activity of the Rho GTPases perturbs the normal development of the eye (17). When overexpressed in transgenic lines, wild type forms of Rho1, Rac1, Rac2, and Cdc42 can each disrupt the normal ommatidial structure of the eye and result in an externally “rough” phenotype (Fig. 3). For example, expression of a Rho1 transgene causes a rough-eye phenotype associated with disruption of the morphology of the photoreceptors (17). In addition, the lattice formed by secondary and tertiary pigment cells as well as the general ommatidial architecture is completely disrupted. Furthermore, disruption of the normal appearance of the polymerized actin in pupal eye discs from these transgenics suggests that the ability of Rho1 to affect actin organization plays an important role in eye morphogenesis. Expression of Rac1, Rac2, and Cdc42 GTPases in the eye similarly disrupts normal eye development, although the observed defects are different in each case, indicating that this system is useful for distinguishing the activities of closely related Rho GTPases in vivo (44).
embryonic ectoderm, and become the imaginal discs during larval development. These discs eventually give rise to all adult tissues. Aside from the commonly observed apical-basal polarity in epithelial tissue culture cells, epithelial cells can also develop a planar polarity relative to the body axis (53). This is reflected in the appearance of distally pointed hairs on the Drosophila wing blades and the regular arrays of trapezoidal shaped rhabdomeres in the Drosophila compound eye. To study the role of Rho GTPases in establishing tissue polarity in the fly wing disc, mutant forms of Rac1 and Cdc42 were expressed specifically in wing discs (54). From those experiments, it was found that Rac1 activity was essential for the proper assembly of cell adherens junctions as well as for the establishment of planar polarity, while Cdc42 was found to be required for epithelial cell shape changes, but is not required for actin assembly at adherens junctions. Neither of these GTPases was required to maintain polarity once it was established. In addition, detailed examination revealed distinct functions for Rac and Cdc42 in regulating different aspects of wing hair formation (55). Specifically, expression of dominant-negative Cdc42 stunts wing hair growth, leading to a failure of wing hair cells to accumulate actin in their distal regions. This suggests that Cdc42 might be responsible for polarized membrane outgrowth which is analogous to the Cdc42 stimulated filopodia formation in fibroblasts. On the other hand, dominant-negative Rac1 promotes the outgrowth of multiple hairs from a single cell, which also exhibit gaps in junctional actin and a disorganized apical microtubule web, indicating that Rac1 is responsible for both actin polymerization and microtubule organization. The existing Rho1 mutant further confirmed the role of Rho GTPases in establishing tissue polarity in wing development (33). Wing clones containing hypomorphic Rho1 alleles exhibit abnormal wing hair polarity. Similarly, in Rho1 eye clones, a large number of ommatidia were incorrectly oriented while the relative position of photoreceptors and accessory cells remain unaltered. This is the same phenotype seen in two previously identified tissue polarity mutants: frizzled and disheveled. Frizzled appears to be a G-protein coupled receptor, and disheveled is a cytoplasmic signaling molecule. Genetic experiments established that Rho1 functions downstream of the frizzled-disheveled mediated tissue polarity pathway (33), which is consistent with the general theme linking G-protein coupled receptors to the Rho signaling pathway.
Tissue Polarity
Summary
Polarized epithelial cells perform a variety of specialized functions and epithelial polarity plays an important role in Drosophila morphogenesis. Drosophila epithelial cells are derived from the invaginated
It has become increasingly clear in the last few years that the Rho family GTPases regulate cytoskeleton rearrangements that are essential for a variety of morphogenetic events associated with the development of
TABLE I
Summary of Rho GTPase Signaling Components Involved in Various Drosophila Developmental Processes Developmental processes Oogenesis: border cell migration Oogenesis: transfer of nurse cell contents to oocytes Gastrulation Dorsal closure
Muscle development Neural development Eye development Tissue polarity: wing development Tissue polarity: eye development
Genes/Pathways involved
References
Rac1
(19)
Rac1, Cdc42, RhoL
(19)
Fog, cta, DRhoGEF2, Rho1 Rho1, Rac1, Cdc42, JNK pathway genes, Dpp pathway genes, and mbc Rac1, mbc Rac1, Cdc42, still life Rho1, Rac1, Rac2, Cdc42 Rac1, Cdc42, Rho1
(24–27)
Rho1, frizzled, disheveled
(31–44)
(18, 43) (18, 49, 50) (17, 44) (33, 54, 55) (33)
possible that DLAR activates Rac1 through a Trio-like GEF to mediate an axon guidance signal. Eye Development
92
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MOLECULAR CELL BIOLOGY RESEARCH COMMUNICATIONS 18b.Luo, L., Liao, Y. J., Jan, L. Y., and Jan, Y. N. (1994) Genes Dev. 8, 1787–1802. 19. Murphy, A. M., and Montell, D. J. (1996) J. Cell Biol. 133, 617– 630. 20. Sasamura, T., Kobayashi, T., Kojima, S., Qadota, H., Ohya, Y., Masai, I., and Hotta, Y. (1997) Mol. Gen. Genet. 254, 486 – 494. 21. Cooley, L., and Theurkauf, W. E. (1994) Science 266, 590 –596. 22. Sweeton, D., Parks, S., Costa, M., and Wieschaus, E. (1991) Development 112, 775–789. 23. Leptin, M. (1995) Annu. Rev. Cell Dev. Biol. 11, 189 –212. 24. Barrett, K., Leptin, M., and Settleman, J. (1997) Cell 91, 905– 915. 25. Hacker, U., and Perrimon, N. (1998) Genes Dev. 12, 274 –284. 26. Costa, M., Wilson, E. T., and Wieschaus, E. (1994) Cell 76, 1075–1089. 27. Parks, S., and Wieschaus, E. (1991) Cell 64, 447– 458. 28. Morize, P., Christiansen, A., Costa, M., Parks, S., and Wieschaus, E. (1998) Dev. Suppl. 125, 589 –597. 29. Gebbink, M. F., Kranenburg, O., Poland, M., van Horck, F. P., Houssa, B., and Moolenaar, W. H. (1997) J. Cell. Biol. 137, 1603–1613. 30. Martinez-Arias, A. (1993) in The Development of Drosophila melanogaster (Bate, M., and Martinez-Arias, A., Eds.), pp. 517– 608. Cold Spring Harbor Laboratory Press, New York, NY. 31. Harden, N., Loh, H. Y., Chia, W., and Lim, L. (1995) Development 121, 903–914. 32. Riesgo-Escovar, J. R., Jenni, M., Fritz, A., and Hafen, E. (1996) Genes Dev. 10, 2759 –2768. 33. Strutt, D. I., Weber, U., and Mlodzik, M. (1997) Nature 387, 292–295. 34. Glise, B., Bourbon, H., and Noselli, S. (1995) Cell 83, 451– 461. 35. Sluss, H. K., Han, Z., Barrett, T., Davis, R. J., and Ip, Y. T. (1996) Genes Dev. 10, 2745–2758. 36. Glise, B., and Noselli, S. (1997) Genes Dev. 11, 1738 –1747. 37. Hou, X. S., Goldstein, E. S., and Perrimon, N. (1997) Genes Dev. 11, 1728 –1737. 38. Kockel, L., Zeitlinger, J., Staszewski, L. M., Mlodzik, M., and Bohmann, D. (1997) Genes Dev. 11, 1748 –1758. 39. Riesgo-Escovar, J. R., and Hafen, E. (1997) Genes Dev. 11, 1717– 1727. 40. Riesgo-Escovar, J. R., and Hafen, E. (1997) Science 278, 669 – 672. 41. Zeitlinger, J., Kockel, L., Peverali, F. A., Jackson, D. B., Mlodzik, M., and Bohmann, D. (1997) EMBO J. 16, 7393–7401. 42. Martin-Blanco, E., Gampel, A., Ring, J., Virdee, K., Kirov, N., Tolkovsky, A. M., and Martinez-Arias, A. (1998) Genes Dev. 12, 557–570. 43. Erickson, M. R., Galletta, B. J., and Abmayr, S. M. (1997) J. Cell Biol. 138, 589 – 603. 44. Nolan, K. M., Barrett, K., Lu, Y., Hu, K. Q., Vincent, S., and Settleman, J. (1998) Genes Dev. 12, 3337–3342. 45. Bate, M. (1993) in The Development of Drosophila melanogaster (Bate, M., and Martinez-Arias, A., Eds.), pp. 1013–1090. Cold Spring Harbor Laboratory Press, New York, NY. 46. Holtzer, H., Strahs, K., Biehl, J., Somlyo, A. P., and Ishikawa, H. (1975) Science 188, 943–945. 47. Hasegawa, H., Kiyokawa, E., Tanaka, S., Nagashima, K., Gotoh, N., Shibuya, M., Kurata, T., and Matsuda, M. (1996) Mol. Cell. Biol. 16, 1770 –1776. 48. Smit, L., van der Horst, G., and Borst, J. (1996) J. Biol. Chem. 271, 8564 – 8569. 49. Sone, M., Hoshino, M., Suzuki, E., Kuroda, S., Kaibuchi, K.,
multicellular organisms. In particular, Drosophila has provided an excellent in vivo system for deciphering the signaling pathways mediated by Rho GTPases, as well as establishing the role of these pathways in numerous developmental processes (Table I). Continued use of this system will undoubtedly lead to the identification of additional Rho signaling components and information regarding the function and organization of the Rho signaling pathways in tissue morphogenesis. The striking similarity between Drosophila and mammalian Rho signaling components identified thus far indicates that the Rho pathways are highly conserved in evolution. Therefore, the findings from the Drosophila system can be extrapolated to higher organisms, including humans. Combined with the rapid progress in the human and Drosophila genome projects, these findings should contribute greatly to our understanding of mammalian Rho GTPase signaling pathways and their roles in normal development and pathological conditions. REFERENCES 1. Van Aelst, L., and D’Souza-Schorey, C. (1997) Genes Dev. 11, 2295–2322. 2. Hall, A. (1998) Science 280, 2074 –2075. 3. Ridley, A. J., and Hall, A. (1992) Cell 70, 389 –399. 4. Ridley, A. J., Paterson, H. F., Johnston, C. L., Diekmann, D., and Hall, A. (1992) Cell 70, 401– 410. 5. Nobes, C. D., and Hall, A. (1995) Cell 81, 53– 62. 6. Watanabe, G., Saito, Y., Madaule, P., Ischizaki, T., Fujisawa, K., Morii, N., Mukai, H., Ono, Y., Kakizuka, S., and Narumiya, S. (1996) Science 271, 645– 648. 7. Amano, M., Mukai, H., Ono, Y., Chihara, K., Matsui, T., Hamajima, Y., Okawa, K., Iwamatsu, A., and Kaibuchi, K. (1996) Science 271, 648 – 650. 8. Quilliam, L. A., Lambert, Q. T., Mickelson-Young, L. A., Westwick, J. K., Sparks, A. B., Kay, B. K., Jenkins, N. A., 9. Gilbert, D. J., Copeland, N. G., and Der, C. J. (1996) J. Biol. Chem. 271, 28772–28776. 10. Vincent, S., and Settleman, J. (1997) Mol. Cell. Biol. 17, 2247– 2256. 11. Leung, T., Chen, X.-Q., Manser, E., and Lim, L. (1996) Mol. Cell. Biol. 16, 5313–5327. 12. Madaule, P., Eda, M., Watanabe, N., Fujisawa, K., Matsuoka, T., Bito, H., Ishizaki, T., and Narumiya, S. (1998) Nature 394, 491– 494. 13. Bagrodia, S., Taylor, S. J., Creasy, C. L., Chernoff, J., and Cerione, R. A. (1995) J. Biol. Chem. 270, 22731–22737. 14. Chong, L. D., Traynor-Kaplan, A., Bokoch, G. M., and Schwartz, M. A. (1994) Cell 79, 507–513. 15. Zhang, J., King, W. G., Dillon, S., Hall, A., Feig, L., and Rittenhouse, S. E. (1993) J. Biol. Chem. 268, 22251–22254. 16. Zheng, Y., Bagrodia, S., and Cerione, R. A. (1994) J. Biol. Chem. 269, 18727–18730. 17. Simon, M. A., Bowtell, D. D. L., Dodson, G. S., Laverty, T. R., and Rubin, G. M. (1991) Cell 67, 701–716. 18a.Hariharan, I. K., Hu, K. Q., Asha, H., Quintanilla, A., Ezzell, R. M., and Settleman, J. (1995) EMBO Journal 14, 292–302. 93
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Nakagoshi, H., Saigo, K., Nabeshima, Y., and Hama, C. (1997) Science 275, 543–547. 50. Kaufmann, N., Wills, Z. P., and Van Vactor, D. (1998) Development 125, 453– 461. 51. Krueger, N. X., Van Vactor, D., Wan, H. I., Gelbart, W. M., Goodman, C. S., and Saito, H. (1996) Cell 84, 611– 622. 52. Debant, A., Serra-Pages, C., Seipel, K., O’Brien, S., Tang, M.,
Park, S. H., and Streuli, M. (1996) Proc. Natl. Acad. Sci. USA 93, 5466 –5471. 53. Eaton, S. (1997) Curr. Opin. Cell Biol. 9, 860 – 866. 54. Eaton, S., Auvinen, P., Luo, L., Jan, Y. N., and Simons, K. (1995) J. Cell Biol. 131, 151–164. 55. Eaton, S., Wepf, R., and Simons, K. (1996) J. Cell Biol. 135, 1277–1289.
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