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Published in final edited form as: Curr Top Dev Biol. 2015 Jan 22;111:375–399. doi: 10.1016/bs.ctdb.2014.11.011

Epithelial Morphogenesis: The Mouse Eye as a Model System1

Bharesh Chauhan *,3, Timothy Plageman †,3, Ming Lou ‡, Richard Lang §,2
PMCID: PMC6014593  NIHMSID: NIHMS947244  PMID: 25662266

Abstract

Morphogenesis is the developmental process by which tissues and organs acquire the shape that is critical to their function. Here, we review recent advances in our understanding of the mechanisms that drive morphogenesis in the developing eye. These investigations have shown that regulation of the actin cytoskeleton is central to shaping the presumptive lens and retinal epithelia that are the major components of the eye. Regulation of the actin cytoskeleton is mediated by Rho family GTPases, by signaling pathways and indirectly, by transcription factors that govern the expression of critical genes. Changes in the actin cytoskeleton can shape cells through the generation of filopodia (that, in the eye, connect adjacent epithelia) or through apical constriction, a process that produces a wedge-shaped cell. We have also learned that one tissue can influence the shape of an adjacent one, probably by direct force transmission, in a process we term inductive morphogenesis. Though these mechanisms of morphogenesis have been identified using the eye as a model system, they are likely to apply broadly where epithelia influence the shape of organs during development.

1. INTRODUCTION

Though the timescales are rather different, biology and geology both define morphogenesis as the study of mechanisms of shape acquisition. Whether the subject is tissue forms or landforms, the goal is to understand the forces of nature that guide the process. For the biologist, the study of morphogenesis is mostly a study of embryonic development (though regeneration is an option). Biologists interested in morphogenesis frequently combine the tools of developmental biology and cell biology. This gives them the means to establish the relationship between the molecular events at the single cell level and the consequence of those events for organismal shape. Though there will be unifying mechanisms of morphogenesis at the cellular scale, the huge complexity of tissue shapes in different species and different organs teaches us that in morphogenesis, details and subtlety matter (an interesting example is the subtle changes in beak shape that allow “Darwin’s finches” to effectively exploit an ecological niche; Mallarino et al., 2011).

Though there are many worthy subjects of study in morphogenesis, in this chapter, we focus on the mouse eye as a model system. Eye morphogenesis in the mouse is very complex and therefore interesting. There are also many tools that can be applied to the study of eye development and so even in these early stages of our understanding, progress has been quite rapid. As a nonessential organ (at least for laboratory mice), the eye also has practical advantages as a subject of study. Finally, perhaps more than any other organ, the shape of the eye is critical for function. When the sensory stimulus is as unforgiving as the photon (just ask anyone over 40 years), the optical characteristics of the tissues and their relative shape and position have to show “inimitable perfection” (Darwin, 1859).

2. EYE MORPHOGENESIS

The basic features of eye morphogenesis that occurs during midgestation in the mouse are shown in Fig. 1. In this chapter, we define two movements of epithelial structures—invagination and evagination—according to the polarity of the epithelium. We define invagination as a coordinated epithelial cell movement in the basal direction and an evagination as an epithelial cell movement in the apical direction. Thus, at E9.5 in the mouse, the optic vesicle (Fig. 1A, ov) is formed through an invagination of the diencephalic neural tube. This invagination brings the distal epithelium of the optic vesicle (the future retina) in close apposition to the embryonic surface ectoderm that will form both the lens and corneal ectoderm. As optic vesicle invagination occurs, the periocular mesenchyme that lies between the optic vesicle and surface ectoderm is excluded. Inductive signaling events (Charlton-Perkins, Brown, & Cook, 2011; Grocott, Tambalo, & Streit, 2012; Gunhaga, 2011; Lang, 2004; Shaham, Menuchin, Farhy, & Ashery-Padan, 2012) result in the formation of the lens placode (Fig. 1B, lp), a thickened domain of the surface ectoderm that will undergo invagination to form the lens pit (Fig. 1C, lpt). Invagination of the lens pit is coordinated with the formation of the optic cup, a bilayered epithelium that ultimately forms the neural retina and retinal pigmented epithelium (RPE). According to our polarity-based definition of epithelial morphogenesis, the presumptive retina undergoes an evagination and the presumptive RPE undergoes an invagination as the optic cup forms. By E11.5 (Fig. 1D), the lens pit has closed to form the lens vesicle (lv) and the surface ectoderm has separated to form the presumptive corneal ectoderm. At this point in development, the basic layout of the major tissues of the eye is established.

Figure 1.

Figure 1

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Drawing describing the basic features of eye morphogenesis. (A) At E9.5 in the mouse, the optic stalk (os) and optic vesicle (ov) have evaginated from the diencephalic neural tube (nt) and approached the overlying surface ectoderm (se). (B) By E9.75, the thickened surface ectoderm of the lens placode (Ip) has formed. (C) By E1.5, the lens pit (pt) has undergone a coordinated invagination with the bilayered optic cup (oc) that incorporates the forming retinal pigmented epithelium. (D) At E11.5, the lens vesicle (lv) has separated from the surface ectoderm and presumptive corneal epithelium (cor).

3. DYNAMIC CHANGES IN THE ACTIN CYTOSKELETON DRIVE MORPHOGENESIS

It is now well accepted that regulation of cell migration and cell shape via modulation of the actin cytoskeleton is central to morphogenesis. Central to modulation of the actin cytoskeleton are the Rho family GTPases, molecular switches that transfer information in the cell and gain specificity through a series of effector proteins with a limited set of downstream molecular targets (Bishop & Hall, 2000). The GTPases exist in active (GTP bound) and inactive (GDP bound) states and are regulated in several ways. The Rho GTPase-activating proteins (Tcherkezian & Lamarche-Vane, 2007) and the guanine nucleotide dissociation inhibitors (Dovas & Couchman, 2005) are negative regulators while the Rho guanine-nucleotide exchange factors (RhoGEFs; Rossman, Der, & Sondek, 2005; Schmidt, Diriong, Mery, Fabbrizio, & Debant, 2002) act positively. These are frequently expressed with tissue-specific expression patterns and can themselves be regulated. The Rho family GTPases can regulate morphology, adhesion, differentiation, proliferation, and vesicle trafficking (Heasman & Ridley, 2008; Linseman & Loucks, 2008; Nobes & Hall, 1994, 1999; Villalonga & Ridley, 2006). In some of the first assessments of Rho family GTPase activity, it was noted that cultured cells produced different types of actin and different cell shapes, depending on which active GTPase was expressed. Cdc42 activation resulted in the formation of filopodia, Rac1 in the formation of lamellipodia and RhoA the formation of contractile actin and stress fibers (Jaffe & Hall, 2005). These findings were important for the developmental biologists interested in morphogenesis, because they provided a starting point for the investigation of these pathways in vivo.

Based on the experimental analysis that will be summarized below, it is clear that a complex series of physical and signaling interactions between the epithelial and mesenchymal components of the developing eye are required to shape a normal structure. The major theme emerging from this review is that even morphogenetic movements that appear very simple are mechanistically quite complex and require input from many pathways. In addition, it is often the case that each pathway contributes just a little to the overall movement. A further level of complexity emerges from the interaction between adjacent tissues. Though we cannot, as a field, claim to have solved the problem of eye morphogenesis, already many mechanisms have been uncovered. What are these mechanisms? Below, we describe what is known, more-or-less in ontological order. Many of the mechanisms so far identified are directly or indirectly linked to modulation of the actin cytoskeleton.

3.1. What is the function of placode formation in epithelial morphogenesis?

The formation of placodes—regions of thickened epithelium—is associated with the morphogenesis of many structures. Extensive studies on placode specification have been reported (Bailey & Streit, 2006; Streit, 2008), but the mechanisms of epithelial thickening have received less attention. The strong association between placode formation and morphogenesis further suggests that this type of epithelial thickening might be a prerequisite for epithelial bending.

In the case of the lens placode that forms between E9.0 and E9.5 in the mouse, there are several mutant lines in which placodal thickening is compromised. These include the germ line Pax6 mutation present in the Small eye mouse (Hill et al., 1991), regulatory mutations that lead to diminished placodal Pax6 expression (Dimanlig, Faber, Auerbach, Makarenkova, & Lang, 2001), as well as conditional deletion of Pax6 in the lens placode (Ashery-Padan, Marquardt, Zhou, & Gruss, 2000). Pax6 has a central role in lens development—it is both necessary (Hill et al., 1991) and sufficient (Altmann, Chow, Lang, & Hemmati-Brivanlou, 1997; Chow, Altmann, Lang, & Hemmati-Brivanlou, 1999) for formation of the lens structure from the placode stage—and so it can be difficult to determine how closely this broadly active transcription factor is associated with the morphogenesis machinery. Despite this, it has been shown, with placodal deletion of Pax6, that there is a loss of expression of Shroom3 (Plageman et al., 2010), an actin scaffold protein with an important role in changing the shape of epithelial cells (Lee, Le, & Wallingford, 2009; Nishimura & Takeichi, 2008; see discussion below). This contrasts with placodal deletion of Sox2, another transcription factor involved in lens development. In the placodal Sox2 deletion, though there are consequences for lens development (Smith, Miller, Radice, Ashery-Padan, & Lang, 2009), Shroom3 expression is retained and placodal invagination initiated (Plageman et al., 2010). Though it is likely that Pax6 regulates many genes after lens induction, these data link the transcription factor to Shroom3 expression and the morphogenesis machinery (Fig. 2).

Figure 2.

Figure 2

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Summary schematic of eye morphogenesis pathways. This schematic describes some of the pathways involved in morphogenesis of the mouse eye from E9.5, when the lens placode overlies the optic vesicle, to E10.5 when the lens pit has invaginated and the optic cup has become bilayered. The surface ectoderm, presumptive retina, and presumptive retina pigmented epithelium are color-coded in red, green, and gray, respectively. Within the red box, we summarize the action of three GTPases that have distinct actions on lens placode invagination. Cdc42 and IRSp53 are required for the generation of filopodia within the presumptive lens that connect to the presumptive retina and permit force transmission between the two epithelia. They are required for a full invagination of the lens pit and for subtle shaping of the optic cup. This is and example where lens pit invagination depends on extrinsic forces. The GTPases RhoA and Rac1 are required for the generation of intrinsic forces for lens pit invagination because they control cell length (Rac1 through an Arpc2/cortactin response) and apical constriction (RhoA through a Rho kinase (ROCK) phospho-myosin pathway). RhoA activity is likely controlled by the TRIO guanine nucleotide exchange factor and in turn, this is required for the apical localization of Shroom3, an actin and ROCK-binding protein required for apical constriction. At the transcriptional level, Shroom3 is positively regulated by Pax6. During lens pit invagination, the coordinated morphogenesis of the optic cup is in part stimulated by Wnt/Catnb pathway ligands that are produced by nonlens surface ectoderm. These ligands stimulate the production of cells within the developing retinal pigmented epithelium (RPE). The RPE thus expands against the relatively static retina and, with the principle of a bimetallic strip, induces curvature in the optic cup. The red arrows indicate the movements of morphogenesis for the each epithelial layer.

Some well-known signaling pathways also contribute to formation of the lens placode. When the fibroblast growth factor (Fgf) signaling response is compromised in the preplacodal ectoderm of the mouse, placodal thickening is reduced (Faber et al., 2001), a result confirmed first by mutation of the Fgf receptor adapter protein Frs2α (Gotoh et al., 2004) and later by conditional Fgf receptor deletion (Garcia et al., 2011). Defective Bmp signaling also results in changes to placode formation. Both Bmp4 and Bmp7 germ-line mutant mice exhibit a failure of lens placode formation (Furuta & Hogan, 1998; Wawersik et al., 1999). The defects in lens morphogenesis and changes in epithelial F-actin distribution that arise when the Bmpr1a and Acvr1 receptors are conditionally deleted postinduction (Rajagopal et al., 2009) suggests that this signaling pathway may be quite closely linked to the morphogenesis machinery. Given existing data (Gamell et al., 2008; Konstantinidis, Moustakas, & Stournaras, 2011; Lee-Hoeflich et al., 2004), it is quite likely that there are several levels at which Bmp signaling can regulate cell shape via modulation of the actin cytoskeleton.

The interaction between the presumptive lens and presumptive retina is important for subsequent morphogenesis. In part, this is because of the inductive signaling that takes place, but is also because a physical association is important for coordinated shape formation in the presumptive lens and retina. The importance of the presumptive lens–retina interface is illustrated by experiments that have examined the role of the extracellular matrix (ECM) and more specifically of fibronectin (FN), in lens morphogenesis. In a placode-conditional Pax6 mutant mouse, ECM deposition between the lens and retina is diminished, suggesting that Pax6 directly or indirectly regulates ECM deposition (Huang et al., 2011). The limited ECM at the presumptive lens–retina interface may contribute to the failure of lens pit invagination in the Pax6 mutant. In addition, when FN1 is conditionally deleted throughout the mouse embryo, there is a failure of both placode thickening and lens pit invagination (Huang et al., 2011). Though this mutant may have many signaling and structural defects, these data suggest that placode thickening may be a prerequisite for invagination. These data are also consistent with descriptive analyses of lens placode thickening, suggesting that an increase in cell density coupled with static cell volumes (Mckeehan, 1951) results in thickening. Additional experimental studies attempting to restrict the outward expansion of presumptive lens placodal cells through physical means resulted in ectopic thickening and/or precocious invagination (Steding, 1967; Wakely, 1984). These data further support the hypothesis that placodal thickening is a prerequisite for invagination. In purely structural terms, this might well make sense: just as a deeper beam can better resist the force of gravity, a thicker epithelium can better generate the forces required for invagination.

3.2. Extrinsic force transmission via filopodia

Events in morphogenesis require precise cell positioning and guidance that can be provided by filopodia. This function can be found in migratory tissue sheets, such as dorsal closure in Drosophila (Martin-Blanco, Pastor-Pareja, & Garcia-Bellido, 2000), ventral closure in Caenorhabditis elegans (Williams-Masson, Malik, & Hardin, 1997), and wound healing (Wood et al., 2002). Filopodia also feature in developing vasculature (Gallo & Letourneau, 2004), neuronal projections (De Smet, Segura, De Bock, Hohensinner, & Carmeliet, 2009), and the Drosophila tracheal system (Wolf, Gerlach, & Schuh, 2002). Studied primarily in fibroblasts and neurite growth cones, filopodia are defined as probing cellular protrusions that aid in migration and extension and that contain bundles of parallel actin filaments (Faix & Rottner, 2006; McClay, 1999; Passey, Pellegrin, & Mellor, 2004; Raich, Agbunag, & Hardin, 1999). Their length differs depending on their tip adhesion; those that are anchored at their tips by cadherins tend to be shorter (Partridge & Marcantonio, 2006; Raich et al., 1999; Wood et al., 2002), whereas the integrin-dependent focal adhesion tips tend to be longer (Partridge & Marcantonio, 2006).

The Rho family GTPase Cdc42 has many functions in the cell, but it is critical for the formation of filopodia (Nobes & Hall, 1995; Ridley, 2006). In vitro, it does so through binding in the active state with the ubiquitously expressed effector protein N-WASP (Banzai, Miki, Yamaguchi, & Takenawa, 2000; Egile et al., 1999; Kurisu & Takenawa, 2009; Miki, Sasaki, Takai, & Takenawa, 1998), a Wiskott–Aldrich syndrome gene-family member. N-WASP subsequently activates the actin nucleator Arp2/3, and as a result, fast growing barbed-end actin filaments extend the plasma membrane to form protrusive filopodia. The formation of Cdc42-stimulated filopodia in N-WASP-deficient fibroblasts revealed an alternative pathway for filopodia formation (Snapper et al., 2001). This was later discovered to require the diaphanous-related formins (DRFs; Peng, Wallar, Flanders, Swiatek, & Alberts, 2003), a group of actin nucleators that function downstream of the Rho GTPases. Furthermore, it has been found that Rif forms filopodia through the action of a DRF member, mDia2. A third filopodiogenesis pathway requires the effector protein IRSp53 interacting with either Mena (Krugmann et al., 2001) or Eps8 (Disanza et al., 2006). IRSp53 and Eps8 are also involved in signal transduction, whereas IRSp53 is a substrate for the insulin receptor (Yeh, Ogawa, Danielsen, & Roth, 1996) and Eps8 is a substrate for other tyrosine kinase receptors (Fazioli et al., 1993).

Investigation of filopodia in morphogenesis began with the secondary mesenchymal cells of the sea urchin Arbacia punctulata and the sand dollar Echinarachniu sparma echinoderm species as early as 1969 (Tilney & Gibbins, 1969). This was later followed by investigation of their contractile function in the tunic cells of the ascidian Botryllus schlosseri, where they were found to change cell shape and regulate cell migration (Izzard, 1974). Here, filopodia were observed to be long (reaching lengths of up to 200 μm). Laser ablation studies of the filopodia arising from secondary mesenchymal cells in the sea urchin suggested that they had a contractile function and aided in generating the forces required for invagination of the archenteron during gastrulation (Hardin, 1988). A role for filopodia in connecting epithelium to mesenchyme was found in the examples of the developing avian lung (Gallagher, 1986), and in the dental mesenchyme making contact with the overlying dental epithelium (Fukumoto et al., 2006). Their role in branching morphogenesis was revealed in avian lung development as well as in Drosophila tracheal development (Samakovlis et al., 1996). Their role in wound healing was found through the series of studies conducted in the cornea (Crosson, Klyce, & Beuerman, 1986; Pfister, 1975), skin (Pang, Daniels, & Buck, 1978; Vasioukhin & Fuchs, 2001), and then in Drosophila dorsal closure (Millard & Martin, 2008; Wood et al., 2002), the latter of which recapitulated a number of embryonic tissue closure mechanisms. The contractile function of filopodia and the specific myosins in the varying filopodia have recently been deciphered (Berg & Cheney, 2002; Chauhan et al., 2009; Jacinto et al., 2002; Moores, Sabry, & Spudich, 1996; Sousa & Cheney, 2005; Tokuo & Ikebe, 2004; Wagner, Barylko, & Albanesi, 1992; Zhang et al., 2004).

Ida Mann’s book on human eye development (Mann, 1950) described “protoplasmic processes” that extended between the epithelia of the presumptive lens and retina. Later, the McAvoy group showed, by TEM, that these processes formed a junction-like structure at the tip where they contacted the presumptive retina (McAvoy, 1980). These structures were shown to be actin-rich and devoid of tubulin, suggesting that they were filopodia (Chauhan et al., 2009). A mutant mouse in which the Rho family GTPase Cdc42 was deleted in the lens placode exhibited an absence of interepithelial filopodia further confirming their identity. This mutant also indicated that most of the filopodia originate in the cells of the lens placode and lens pit. A series of mutant mice, including the Cdc42 placode conditional mutant, a germ line mutant for the Cdc42 effector IRSp53, and a conditional mutant for focal adhesion kinase, revealed a spectrum of filopodial loss and an inverse correlation with interepithelial distance. In the Cdc42 conditional mutant where all filopodia were lost, about one-third of the lens pit invagination depth was also lost, suggesting that these structures served to transmit forces between presumptive retina and lens as a means of refining the process of morphogenesis. This suggestion was consistent with the observation that these filopodia contained active phosphomyosin II and were actively contractile. Filopodia-deficient mutants, not surprisingly, show changes in the shape of the lens pit. However, they also show changes in the shape of the adjacent retinal cup (Fig. 3), the structure to which they connect, probably via integrin-mediated adhesion. These nonautonomous epithelial shape changes provide some evidence that one of the functions of these filopodia is to transmit forces from one epithelium to another. Drawing a parallel with the classical concept of inductive fate change, we refer to this phenomenon as inductive morphogenesis. This analysis indicates that the forces required to drive morphogenesis can be extrinsic (Fig. 2).

Figure 3.

Figure 3

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Inductive morphogenesis in the epithelia of the developing eye. (A) The average shape of the basal surfaces of the lens pit and presumptive retina in a wild-type, E10.5 mouse eye. The black line represents that average shape and the green (gray in the print version) shaded area, two standard deviations from the average shape. (B) The average shape of the basal surfaces of the lens pit and presumptive retina in an Le-cre; Cdc42fl/fl, E10.5 mouse eye. In this mutant (Cdc42 M), there are no filopodia connecting the epithelia. As a consequence, the interepithelial distance has increased. The lens pit (C) has lost about one-third of the invagination distance and the retinal cup (D) is also flatter. Since genetic changes in this mouse line are restricted to the presumptive lens, this provides some evidence for force transmission between the epithelia. These data were generated as described previously (Chauhan et al., 2009).

3.3. Intrinsic force generation through apical constriction and cell elongation

When an epithelium invaginates, the cells within the epithelium must, on average, adopt a wedge-shaped morphology. There is now much evidence at both the cellular and molecular levels that this shape change is primarily a result of a reduced apical area that is the result of a process referred to as apical constriction (AC). For the transition between the lens placode and lens pit (Fig. 1) an analysis of cell shape showed that both apical constriction and cell elongation accompanied invagination (Hendrix & Zwaan, 1974). Below, we describe some of the molecular mediators of and epithelial cell elongation and AC and indicate how their activities are integrated.

3.3.1 Apical constriction

Many of the significant advances in our understanding of AC mechanisms have come from analyzing invertebrate morphogenesis including Drosophila gastrulation and dorsal closure. During gastrulation, coordinated AC along the ventral embryonic axis drives mesodermal invagination (Leptin, 2005). Mesodermal cells undergo AC through the contraction of an apically positioned network of actomyosin that spans the apical cortex (Dawes-Hoang et al., 2005; Martin, Kaschube, & Wieschaus, 2009). Assembly and contraction of myosin filaments are facilitated through the phosphorylation and activation of myosin regulatory light chain and phosphorylation and deactivation of myosin phosphatases (Bresnick, 1999). The pathway leading to these phosphorylation events in Drosophila gastrulation are triggered by a well-understood cascade of events that commences with the mesodermal-specific expression of G protein-coupled receptors that apically recruit a guanine exchange factor (DRhoGEF2). DRhoGEF2 activates the Rho-GTPase, Rho1 and in turn stimulates phosphorylation by activating the serine-threonine kinase Rok (Sawyer et al., 2010). Once active, contraction occurs in a step-wise manner consisting of pulses interspersed by periods of relaxation or apical surface stabilization (Azevedo et al., 2011; Blanchard, Murugesu, Adams, Martinez-Arias, & Gorfinkiel, 2010; David, Tishkina, & Harris, 2010; Martin et al., 2009). A molecular connection bridging actomyosin to junctional complexes are also necessary to draw the apical circumference inward and when disrupted the degree of AC is attenuated (Dawes-Hoang et al., 2005; Roh-Johnson et al., 2012; Sawyer, Harris, Slep, Gaul, & Peifer, 2009).

AC within cells of the lens placode is remarkably similar to that observed in Drosophila gastrulation. During lens placode invagination, the vertebrate equivalents of the RhoGEF2-Rho1-Rok pathway of myosin activation are significant contributors. Pharmacological studies inhibiting Rock1/2, the vertebrate homolog of Rok, myosin, or actin dynamics all disrupt lens invagination presumably through the inhibition of AC (Borges, Lamers, Forti, Santos, & Yan, 2011; Plageman et al., 2011). Genetic deletion of RhoA or the RhoA-GEF Trio, functional counterparts of RhoA, and DRhoGEF2, respectively, inhibits AC in the lens and decreased apical levels of activated myosin, indicating a strong molecular conservation of this pathway between species (Chauhan, Lou, Zheng, & Lang, 2011; Plageman et al., 2011). Shroom3, a Rock1/2 and actin-binding cytoskeletal protein, initially identified as a protein required for neural tube AC and morphogenesis, is also necessary for lens placode AC (Haigo, Hildebrand, Harland, & Wallingford, 2003; Hildebrand & Soriano, 1999; Plageman et al., 2010).

Shroom3 appears to play a central role in AC during lens placode invagination. First, its expression domain is regionally restricted to the cells which are undergoing AC. This is likely due in part to lens placode-specific transcription factors, many of which have restricted expression, an idea supported by the observation that Shroom3 expression is dependent upon Pax6 (Plageman et al., 2010; Fig. 2). Second, Shroom3 is sufficient to induce AC and facilitate the assembly of myosin into sarcomeric-like structures in the apical junctions of cultured epithelial cells (Hildebrand, 2005). Third, the absence of Shroom3 causes a reduction in the apical distribution of both F-actin and myosin in the lens placode (Plageman et al., 2010). Its ability to recruit F-actin is likely dependent on the Mena-family of actin modulators which themselves play a role in AC (Plageman et al., 2010; Roffers-Agarwal, Xanthos, Kragtorp, & Miller, 2008). Given that the function of Shroom3 was also shown to be dependent on RhoA, Rock1, and myosin activity, we posit that Shroom3 acts as a nexus of AC activity in the lens by organizing and/or recruiting most if not all of the necessary components.

Like invertebrate AC, the association of actomyosin with the apical junctions is essential for lens placode AC but unlike Drosophila mesodermal cells, much of the actomyosin in the lens placode is found in the circumferential belt positioned near the apical junctional complex (Lang, Herman, Reynolds, Hildebrand, & Plageman, 2014). In a somewhat surprising degree of conservation, actomyosin filaments spanning the apical cortex are indeed present in a subset of lens placodal cells and appear to immediately precede AC. Like invertebrate actomyosin filaments, those in lens placode epithelial cells are connected to deformations in the bicellular junctions (Lang et al., 2014) suggesting that they are contractile. Bicellular deformations and AC are both prevented by the myosin inhibitor blebbistatin (Lang et al., 2014). Further analysis demonstrated that the cadherin-binding modulator of Rho-GTPases, p120-catenin, is required for this junctional displacement and AC in the lens (Lang et al., 2014). This suggests that p120 may be playing a significant role in mediating the connection of actomyosin filaments and/or the circumferential actomyosin belt to the apical junctions. In turn, these mediators are likely critical for the generation of intrinsic forces that can bend an epithelium (Fig. 2).

3.3.2 Cell elongation

In cultured cells, the Rac1 GTPase stimulates the formation of lamellipodia (Jaffe & Hall, 2005). This unique type of cellular process is a consequence of the particular effector molecules that this GTPase activates and in turn, the type of actin complexes that are formed (Jaffe & Hall, 2005). One well-known Rac1 effector is the Wiskott–Aldrich syndrome protein family verprolin homolog WAVE (Kurisu & Takenawa, 2009; Miki, Suetsugu, & Takenawa, 1998) which, through upstream activation of the p21-activated kinase (PAK) effector and binding to the actin nucleator Arp2/3, encourages polymerization of the Y-branched actin networks that give lamellipodia and membrane ruffles (Burridge & Wennerberg, 2004). It has also been shown that active PAK1 has an additional role in membrane transport of WAVE2 to the leading edge for lamellipodia formation (Takahashi & Suzuki, 2009). Interestingly, it has been shown using fluorescence resonance energy transfer microscopy in cultured cells that RacGTP levels are the highest at the leading edge of a migrating cell (Kraynov et al., 2000), and that integrin–matrix interactions may play a role in localized Rac1 activity.

When Rac1 is conditionally deleted from the lens placode, the constituent cells exhibit reduced length (Chauhan et al., 2011). This is likely to be the in vivo manifestation, in the epithelial cell type, of the activity that forms lamellapodia in migrating cultured cells. From the point of view of tissue structure, it may be that this type of cell elongation is important for epithelial bending in that it extends the length of the levers that can be used to generate force.

3.3.3 Rac1, RhoA pathways integrate to control cell shape in the lens placode

It is clear that in some settings, the Rac1 and RhoA actin modulation pathways are mutually antagonistic (Machacek et al., 2009; Xu et al., 2003). In the invaginating lens placode the basic phenotypic response, where lens pit curvature is reduced in RhoA mutants and increased in Rac1 mutants (Chauhan et al., 2011), also suggested this might be the case in vivo. An assessment of cell shapes further showed that, as would be expected, RhoA was required for AC (Chauhan et al., 2011). However, in addition, RhoA mutant lens placode cells also showed increased length. In a reciprocal set of responses, Rac1 mutant placode cells showed reduced length, as would be expected, but also showed increased AC (Chauhan et al., 2011). How can these findings be explained mechanistically?

Using independent markers of the RhoA and Rac1 pathway responses, it was shown that in the RhoA mutant lens placode, there were indications of Rac1 pathway gain of function. For example, the Arpc2 actin nucleation complex subunit was significantly over-represented basally in the placodal RhoA mutant. This could explain why a RhoA mutant epithelial cell had become longer than normal. Similarly, in Rac1 mutant placode cells, phosphomyosin was over-represented at the cell apex. This suggested that the Rac1 mutant placodal cell had a RhoA pathway gain of function and was an explanation for the increased AC and the tighter curvature of the lens pit in this mutant. These observations suggest that during lens placode invagination, RhoA and Rac1 are mutually antagonistic. Furthermore, the data suggest their activities are balanced in a way that gives fine control of cell height and apical width (Chauhan et al., 2011). Since, the ratio of these two cellular dimensions controls the angle that an epithelial cell occupies, in aggregate, Rac1 and RhoA mutual antagonism regulates epithelial curvature (Fig. 2).

There are many examples of a dramatic change in epithelial curvature over just a few cell diameters, and so it is also possible that the Rac1 to RhoA activity ratio can also be controlled locally, presumably by the signaling responses known to regulate these GTPases (Raftopoulou & Hall, 2004). These are likely to be general mechanisms for regulating epithelial morphogenesis given the ubiquity of the Rho family GTPases. Studies have shown the mechanism of RhoA–Rac1 mutual antagonism can be mediated through either p190Rho-GAP (Nimnual, Taylor, & Bar-Sagi, 2003), the kinase sticky (D’Avino, Savoian, & Glover, 2004), or FilGAP (Ohta, Hartwig, & Stossel, 2006). It remains to be determined whether these mechanisms apply to morphogenesis of the eye.

4. SHAPING OF THE OPTIC CUP

Analysis of the role of GTPases and the AC machinery in the invaginating lens pit has made a strong case that the forces that drive epithelial bending are generated intrinsically through cell shape change and extrinsically, via filopodia that connect the invaginating lens placode to the presumptive retina. This arrangement raises the question of whether there is morphogenetic induction—a change in shape dependent on the shape of the adjacent epithelium—that is polarized or reciprocal.

4.1. Does the optic cup pull on the invaginating lens?

Several lines of evidence indicate that the optic cup influences the shape of the lens pit. In several lines of mutant mice, where interepithelial filpodia are reduced in number, there is a reciprocal relationship between the number of filopodia and interepithelial distance. In other words, as the number of filopodia reduces, so does the depth of lens pit invagination. This suggests quite strongly that the optic cup pulls on the lens pit. There is some subtlety, however, in that even in a mouse in which all filopodia are absent, only about one-third of the lens pit invagination depth is lost (Chauhan et al., 2009). This is consistent with a model in which the lens placode has intrinsic force generation (through cell shape change) and that the extrinsic force generation through connecting filopodia is a fine-tuning mechanism (Fig. 2).

4.2. Does the invaginating lens influence the shape of the optic cup?

Recent experiments in which optic cups are developed in culture from ES cells have suggested that the optic cup develops autonomously and is self-shaping (Eiraku et al., 2011). Though the reality is more complicated (see below), the observation that the optic cups forms in the absence of lens has suggested that the invaginating lens pit does not push on the presumptive retina as the optic cup forms. To extend this line of questioning further, we can ask whether there are any changes in the shape of the optic cup when the invaginating lens placode and presumptive retina are uncoupled. Arguably, an uncoupling occurs in Cdc42 conditional mutant mice in which the presumptive lens does not generate any connecting filopodia. When we analyze the shape of the optic cup in this mutant, we find that it is changed compared to a control (Fig. 3). Obviously, this suggests that the invaginating lens placode does influence the shape of the optic cup. As in the reciprocal interaction, the changes in the shape are subtle and so again it might be argued that the lens finetunes the shape of the cup. It is also worth noting that when optic cups are generated in culture, very few of them are perfect in shape and a proportion never undergo the evagination of the presumptive retina that generates the bilayered optic cup (Eiraku et al., 2011; Nakano et al., 2012). This observation contrasts with the in vivo development of the eye where the structure is perfect nearly all of the time. This contrast suggests that while the optic cup can form its basic shape autonomously some of the time, if it is to do so all the time, the influence of surrounding tissues is required. Below, we review the evidence that the surface ectoderm of the eye produces signals that are critical for morphogenesis of the optic cup.

4.3. The “bimetallic strip” mechanism of optic cup morphogenesis

In the mouse, when the lens placode and adjacent ectoderm are prevented from producing Wnt ligands, there are profound consequences for eye development. One consequence is that the optic cup does not form its normal, near-spherical shape but is flat and saucer shaped (Carpenter et al., 2014). This contrasts with the hypothesis that the optic cup is self-shaping and shows very directly that signaling ligands produced by the ectoderm influence optic cup morphogenesis. With the available data, the most compelling hypothesis to explain this function of Wnt ligands is that they are produced by the surface ectoderm and stimulate a Wnt/β-catenin response in the presumptive RPE adjacent to the retinal rim (Carpenter et al., 2014). It is also understood that the Wnt/β-catenin response upregulates the retinoic acid response pathway, known to be critical for optic cup morphogenesis in the RPE. In turn, the Wnt/β-catenin and retinoic acid responses stimulate the production of cells within the RPE that helps to shape the optic cup.

It has been suggested that the mechanism by which RPE cells shape the optic cup is, in principle, similar to the mechanism by which a bimetallic strip bends when it is heated. A bimetallic strip is a temperature sensing device that consists of two different metals bonded along their length. If these metals have different expansion coefficients, then the bimetallic strip will bend when it is heated (https://www.youtube.com/watch?v=sP5NwEkd3ds). In the same way, as more RPE cells are produced in the bilayered optic cup, they will induce curvature (Fig. 2).

The bimetallic strip model of optic cup morphogenesis is consistent with a number of published observations. First, it has been shown that the presumptive RPE is Wnt/β-catenin responsive according to the expression of Wnt/β-catenin pathway components and reporters (Cho & Cepko, 2006; Fuhrmann, Levine, & Reh, 2000; Liu, Mohamed, Dufort, & Wallace, 2003; Liu, Thurig, Mohamed, Dufort, & Wallace, 2006; Liu et al., 2007; Trimarchi, Cho, & Cepko, 2009) and the expression of Wnt/β-catenin target genes (Eiraku et al., 2011; Fuhrmann, 2008; Westenskow, Piccolo, & Fuhrmann, 2009). Second, assessment of the stiffness of the ocular epithelia using the atomic force microscope has indicated that while the RPE is stiff, the presumptive retina is not (Eiraku et al., 2011). This suggests, as does the bimetallic strip model that the RPE acts as a scaffold and is perhaps the dominant structural determinant of optic cup curvature (Fig. 2).

How do we reconcile the apparently conflicting observations that in vivo, optic morphogenesis is dependent on Wnt ligands from the surface ectoderm while in culture, they can form in the absence of the surface ectoderm? The simple reconciliation is that in the ES-derived optic cups, these critical natural signaling ligands are provided as media supplements. Retinoic acid is a standard component of “optic cup medium” and supplementary Wnt ligands have been shown to enhance the developmental process (Eiraku & Sasai, 2012).

5. CONCLUSIONS

A defining message of the current studies in eye morphogenesis is that many small influences can lead to dramatic shape changes when each of those subtle influences are focused on a single structure. The eye has a structure that is more complex than most organs, and so it will be interesting to discover if this principle holds elsewhere.

What does the future hold for studies of morphogenesis, including those focused on the eye? As the field progresses, it will be important to quantify morphogenesis numerically and, perhaps using force sensors (Grashoff et al., 2010), define the forces, in real time, that drive morphogenesis. This, coupled to a more sophisticated understanding of the signaling pathways that drive the morphogenesis machinery, will steadily build a comprehensive model of the process.

Acknowledgments

We would especially like to thank Dr. Sonya Faber for the drawing of eye morphogenesis shown in Fig. 1. This work was supported by Grant R01 EY016241 from the National Eye Institute of the National Institutes of Health and by funds from the Abrahamson Pediatric Eye Institute of Cincinnati Children’s Hospital.

Footnotes

1

This chapter is dedicated to the memory of Yoshiki Sasai, an exceptional scientist whose work contributed critically to many fields of enquiry, including the one reviewed here.

References

  1. Altmann CR, Chow RL, Lang RA, Hemmati-Brivanlou A. Lens induction by Pax-6 in Xenopus laevis. Developmental Biology. 1997;185(1):119–123. doi: 10.1006/dbio.1997.8573. http://dx.doi.org/10.1006/dbio.1997.8573. [DOI] [PubMed] [Google Scholar]
  2. Ashery-Padan R, Marquardt T, Zhou X, Gruss P. Pax6 activity in the lens primordium is required for lens formation and for correct placement of a single retina in the eye. Genes & Development. 2000;14(21):2701–2711. doi: 10.1101/gad.184000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Azevedo D, Antunes M, Prag S, Ma X, Hacker U, Brodland GW, et al. DRhoGEF2 regulates cellular tension and cell pulsations in the Amnioserosa during Drosophila dorsal closure. PLoS One. 2011;6(9):e23964. doi: 10.1371/journal.pone.0023964. http://dx.doi.org/10.1371/journal.pone.0023964. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bailey AP, Streit A. Sensory organs: Making and breaking the pre-placodal region. Current Topics in Developmental Biology. 2006;72:167–204. doi: 10.1016/S0070-2153(05)72003-2. http://dx.doi.org/10.1016/S0070-2153(05)72003-2. [DOI] [PubMed] [Google Scholar]
  5. Banzai Y, Miki H, Yamaguchi H, Takenawa T. Essential role of neural Wiskott-Aldrich syndrome protein in neurite extension in PC12 cells and rat hippocampal primary culture cells. Journal of Biological Chemistry. 2000;275(16):11987–11992. doi: 10.1074/jbc.275.16.11987. [DOI] [PubMed] [Google Scholar]
  6. Berg JS, Cheney RE. Myosin-X is an unconventional myosin that undergoes intrafilopodial motility. Nature Cell Biology. 2002;4(3):246–250. doi: 10.1038/ncb762. http://dx.doi.org/10.1038/ncb762. [DOI] [PubMed] [Google Scholar]
  7. Bishop AL, Hall A. Rho GTPases and their effector proteins. Biochemical Journal. 2000;348(Pt 2):241–255. [PMC free article] [PubMed] [Google Scholar]
  8. Blanchard GB, Murugesu S, Adams RJ, Martinez-Arias A, Gorfinkiel N. Cytoskeletal dynamics and supracellular organisation of cell shape fluctuations during dorsal closure. Development. 2010;137(16):2743–2752. doi: 10.1242/dev.045872. http://dx.doi.org/10.1242/dev.045872. [DOI] [PubMed] [Google Scholar]
  9. Borges RM, Lamers ML, Forti FL, Santos MF, Yan CY. Rho signaling pathway and apical constriction in the early lens placode. Genesis. 2011;49(5):368–379. doi: 10.1002/dvg.20723. http://dx.doi.org/10.1002/dvg.20723. [DOI] [PubMed] [Google Scholar]
  10. Bresnick AR. Molecular mechanisms of nonmuscle myosin-II regulation. Current Opinion in Cell Biology. 1999;11(1):26–33. doi: 10.1016/s0955-0674(99)80004-0. [DOI] [PubMed] [Google Scholar]
  11. Burridge K, Wennerberg K. Rho and Rac take center stage. Cell. 2004;116(2):167–179. doi: 10.1016/s0092-8674(04)00003-0. [DOI] [PubMed] [Google Scholar]
  12. Carpenter AC, Smith AN, Wagner H, Wallace VA, Ashery-Padan R, Lang RA. Wnt ligands from the embryonic surface ectoderm regulate “bimetallic strip” optic cup morphogenesis in the mouse. Development. 2014 doi: 10.1242/dev.120022. in press. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Charlton-Perkins M, Brown NL, Cook TA. The lens in focus: A comparison of lens development in Drosophila and vertebrates. Molecular Genetics & Genomics. 2011;286(3-4):189–213. doi: 10.1007/s00438-011-0643-y. http://dx.doi.org/10.1007/s00438-011-0643-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Chauhan BK, Disanza A, Choi SY, Faber SC, Lou M, Beggs HE. Cdc42- and IRSp53-dependent contractile filopodia tether presumptive lens and retina to coordinate epithelial invagination. Development. 2009;136(21):3657–3667. doi: 10.1242/dev.042242. http://dx.doi.org/10.1242/dev.042242. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Chauhan BK, Lou M, Zheng Y, Lang RA. Balanced Rac1 and RhoA activities regulate cell shape and drive invagination morphogenesis in epithelia. Proceedings of the National Academy of Sciences of the United States of America. 2011;108(45):18289–18294. doi: 10.1073/pnas.1108993108. http://dx.doi.org/10.1073/pnas.1108993108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Cho SH, Cepko CL. Wnt2b/beta-catenin-mediated canonical Wnt signaling determines the peripheral fates of the chick eye. Development. 2006;133(16):3167–3177. doi: 10.1242/dev.02474. [DOI] [PubMed] [Google Scholar]
  17. Chow RL, Altmann CR, Lang RA, Hemmati-Brivanlou A. Pax6 induces ectopic eyes in a vertebrate. Development. 1999;126(19):4213–4222. doi: 10.1242/dev.126.19.4213. [DOI] [PubMed] [Google Scholar]
  18. Crosson CE, Klyce SD, Beuerman RW. Epithelial wound closure in the rabbit cornea. A biphasic process. Investigative Ophthalmology & Visual Science. 1986;27(4):464–473. [PubMed] [Google Scholar]
  19. D’A vino PP, Savoian MS, Glover DM. Mutations in sticky lead to defective organization of the contractile ring during cytokinesis and are enhanced by Rho and suppressed by Rac. Journal of Cell Biology. 2004;166(1):61–71. doi: 10.1083/jcb.200402157. http://dx.doi.org/10.1083/jcb.200402157. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Darwin C. The origin of species. London: John Murray; 1859. [Google Scholar]
  21. David DJ, Tishkina A, Harris TJ. The PAR complex regulates pulsed actomyosin contractions during amnioserosa apical constriction in Drosophila. Development. 2010;137(10):1645–1655. doi: 10.1242/dev.044107. http://dx.doi.org/10.1242/dev.044107. [DOI] [PubMed] [Google Scholar]
  22. Dawes-Hoang RE, Parmar KM, Christiansen AE, Phelps CB, Brand AH, Wieschaus EF. folded gastrulation, cell shape change and the control of myosin localization. Development. 2005;132(18):4165–4178. doi: 10.1242/dev.01938. [DOI] [PubMed] [Google Scholar]
  23. De Smet F, Segura I, De Bock K, Hohensinner PJ, Carmeliet P. Mechanisms of vessel branching: Filopodia on endothelial tip cells lead the way. Arteriosclerosis, Thrombosis, and Vascular Biology. 2009;29(5):639–649. doi: 10.1161/ATVBAHA.109.185165. http://dx.doi.org/10.1161/ATVBAHA.109.185165. [DOI] [PubMed] [Google Scholar]
  24. Dimanlig PV, Faber SC, Auerbach W, Makarenkova HP, Lang RA. The upstream ectoderm enhancer in Pax6 has an important role in lens induction. Development. 2001;128(22):4415–4424. doi: 10.1242/dev.128.22.4415. [DOI] [PubMed] [Google Scholar]
  25. Disanza A, Mantoani S, Hertzog M, Gerboth S, Frittoli E, Steffen A, et al. Regulation of cell shape by Cdc42 is mediated by the synergic actin-bundling activity of the Eps8–IRSp53 complex. Nature Cell Biology. 2006;8(12):1337–1347. doi: 10.1038/ncb1502. [DOI] [PubMed] [Google Scholar]
  26. Dovas A, Couchman JR. RhoGDI: Multiple functions in the regulation of Rho family GTPase activities. Biochemical Journal. 2005;390(Pt 1):1–9. doi: 10.1042/BJ20050104. http://dx.doi.org/10.1042/BJ20050104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Egile C, Loisel TP, Laurent V, Li R, Pantaloni D, Sansonetti PJ, et al. Activation of the CDC42 effector N-WASP by the Shigella flexneri IcsA protein promotes actin nucleation by Arp2/3 complex and bacterial actin-based motility. Journal of Cell Biology. 1999;146(6):1319–1332. doi: 10.1083/jcb.146.6.1319. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Eiraku M, Sasai Y. Mouse embryonic stem cell culture for generation of three-dimensional retinal and cortical tissues. Nature Protocols. 2012;7(1):69–79. doi: 10.1038/nprot.2011.429. http://dx.doi.org/10.1038/nprot.2011.429. [DOI] [PubMed] [Google Scholar]
  29. Eiraku M, Takata N, Ishibashi H, Kawada M, Sakakura E, Okuda S. Selforganizing optic-cup morphogenesis in three-dimensional culture. Nature. 2011;472(7341):51–56. doi: 10.1038/nature09941. http://dx.doi.org/10.1038/nature09941. [DOI] [PubMed] [Google Scholar]
  30. Faber SC, Dimanlig P, Makarenkova HP, Shirke S, Ko K, Lang RA. Fgf receptor signaling plays a role in lens induction. Development. 2001;128(22):4425–4438. doi: 10.1242/dev.128.22.4425. [DOI] [PubMed] [Google Scholar]
  31. Faix J, Rottner K. The making of filopodia. Current Opinion in Cell Biology. 2006;18(1):18–25. doi: 10.1016/j.ceb.2005.11.002. http://dx.doi.org/10.1016/jxeb.2005.11.002. [DOI] [PubMed] [Google Scholar]
  32. Fazioli F, Minichiello L, Matoska V, Castagnino P, Miki T, Wong WT, et al. Eps8, a substrate for the epidermal growth factor receptor kinase, enhances EGF-dependent mitogenic signals. The EMBO Journal. 1993;12(10):3799–3808. doi: 10.1002/j.1460-2075.1993.tb06058.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Fuhrmann S. Wnt signaling in eye organogenesis. Organogenesis. 2008;4(2):60–67. doi: 10.4161/org.4.2.5850. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Fuhrmann S, Levine EM, Reh TA. Extraocular mesenchyme patterns the optic vesicle during early eye development in the embryonic chick. Development. 2000;127(21):4599–4609. doi: 10.1242/dev.127.21.4599. [DOI] [PubMed] [Google Scholar]
  35. Fukumoto S, Miner JH, Ida H, Fukumoto E, Yuasa K, Miyazaki H, et al. Laminin alpha5 is required for dental epithelium growth and polarity and the development of tooth bud and shape. Journal of Biological Chemistry. 2006;281(8):5008–5016. doi: 10.1074/jbc.M509295200. http://dx.doi.org/10.1074/jbc.M509295200. [DOI] [PubMed] [Google Scholar]
  36. Furuta Y, Hogan BLM. BMP4 is essential for lens induction in the mouse embryo. Genes & Development. 1998;12(23):3764–3775. doi: 10.1101/gad.12.23.3764. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Gallagher BC. Branching morphogenesis in the avian lung: Electron microscopic studies using cationic dyes. Journal of Embryology and Experimental Morphology. 1986;94:189–205. [PubMed] [Google Scholar]
  38. Gallo G, Letourneau PC. Regulation of growth cone actin filaments by guidance cues. Journal of Neurobiology. 2004;58(1):92–102. doi: 10.1002/neu.10282. http://dx.doi.org/10.1002/neu.10282. [DOI] [PubMed] [Google Scholar]
  39. Gamell C, Osses N, Bartrons R, Ruckle T, Camps M, Rosa JL, et al. BMP2 induction of actin cytoskeleton reorganization and cell migration requires PI3-kinase and Cdc42 activity. Journal of Cell Science. 2008;121(Pt. 23):3960–3970. doi: 10.1242/jcs.031286. http://dx.doi.org/10.1242/jcs.031286. [DOI] [PubMed] [Google Scholar]
  40. Garcia CM, Huang J, Madakashira BP, Liu Y, Rajagopal R, Dattilo L, et al. The function of FGF signaling in the lens placode. Developmental Biology. 2011;351(1):176–185. doi: 10.1016/j.ydbio.2011.01.001. http://dx.doi.org/10.1016/j.ydbio.2011.01.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Gotoh N, Ito M, Yamamoto S, Yoshino I, Song N, Wang Y. Tyrosine phosphorylation sites on FRS2alpha responsible for Shp2 recruitment are critical for induction of lens and retina. Proceedings of the National Academy of Sciences of the United States of America. 2004;101(49):17144–17149. doi: 10.1073/pnas.0407577101. http://dx.doi.org/10.1073/pnas.0407577101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Grashoff C, Hoffman BD, Brenner MD, Zhou R, Parsons M, Yang MT, et al. Measuring mechanical tension across vinculin reveals regulation of focal adhesion dynamics. Nature. 2010;466(7303):263–266. doi: 10.1038/nature09198. http://dx.doi.org/10.1038/nature09198. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Grocott T, Tambalo M, Streit A. The peripheral sensory nervous system in the vertebrate head: A gene regulatory perspective. Developmental Biology. 2012;370(1):3–23. doi: 10.1016/j.ydbio.2012.06.028. http://dx.doi.org/10.1016/j.ydbio.2012.06.028. [DOI] [PubMed] [Google Scholar]
  44. Gunhaga L. The lens: A classical model of embryonic induction providing new insights into cell determination in early development. Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences. 2011;366(1568):1193–1203. doi: 10.1098/rstb.2010.0175. http://dx.doi.org/10.1098/rstb.2010.0175. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Haigo SL, Hildebrand JD, Harland RM, Wallingford JB. Shroom induces apical constriction and is required for hingepoint formation during neural tube closure. Current Biology. 2003;13(24):2125–2137. doi: 10.1016/j.cub.2003.11.054. [DOI] [PubMed] [Google Scholar]
  46. Hardin J. The role of secondary mesenchyme cells during sea urchin gastrulation studied by laser ablation. Development. 1988;103(2):317–324. doi: 10.1242/dev.103.2.317. [DOI] [PubMed] [Google Scholar]
  47. Heasman SJ, Ridley AJ. Mammalian Rho GTPases: New insights into their functions from in vivo studies. Nature Reviews Molecular Cell Biology. 2008;9(9):690–701. doi: 10.1038/nrm2476. http://dx.doi.org/10.1038/nrm2476. [DOI] [PubMed] [Google Scholar]
  48. Hendrix RW, Zwaan J. Cell shape regulation and cell cycle in embryonic lens cells. Nature. 1974;247(437):145–147. doi: 10.1038/247145a0. [DOI] [PubMed] [Google Scholar]
  49. Hildebrand JD. Shroom regulates epithelial cell shape via the apical positioning of an actomyosin network. Journal of Cell Science. 2005;118(Pt. 22):5191–5203. doi: 10.1242/jcs.02626. [DOI] [PubMed] [Google Scholar]
  50. Hildebrand JD, Soriano P. Shroom, a PDZ domain-containing actin-binding protein, is required for neural tube morphogenesis in mice. Cell. 1999;99(5):485–497. doi: 10.1016/s0092-8674(00)81537-8. [DOI] [PubMed] [Google Scholar]
  51. Hill RE, Favor J, Hogan BL, Ton CC, Saunders GF, Hanson IM, et al. Mouse small eye results from mutations in a paired-like homeobox-containing gene. Nature. 1991;354(6354):522–525. doi: 10.1038/354522a0. [DOI] [PubMed] [Google Scholar]
  52. Huang J, Rajagopal R, Liu Y, Dattilo LK, Shaham O, Ashery-Padan R, et al. The mechanism of lens placode formation: A case of matrix-mediated morphogenesis. Developmental Biology. 2011;355(1):32–42. doi: 10.1016/j.ydbio.2011.04.008. http://dx.doi.org/10.1016/j.ydbio.2011.04.008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Izzard CS. Contractile filopodia andin vivo cell movement in the tunic of the ascidian, Botryllus schlossen. Journal of Cell Science. 1974;15(3):513–535. doi: 10.1242/jcs.15.3.513. [DOI] [PubMed] [Google Scholar]
  54. Jacinto A, Wood W, Woolner S, Hiley C, Turner L, Wilson C, et al. Dynamic analysis of actin cable function during Drosophila dorsal closure. Current Biology. 2002;12(14):1245–1250. doi: 10.1016/s0960-9822(02)00955-7. [DOI] [PubMed] [Google Scholar]
  55. Jaffe AB, Hall A. Rho GTPases: Biochemistry and biology. Annual Review of Cell and Developmental Biology. 2005;21:247–269. doi: 10.1146/annurev.cellbio.21.020604.150721. [DOI] [PubMed] [Google Scholar]
  56. Konstantinidis G, Moustakas A, Stournaras C. Regulation of myosin light chain function by BMP signaling controls actin cytoskeleton remodeling. Cellular Physiology and Biochemistry. 2011;28(5):1031–1044. doi: 10.1159/000335790. http://dx.doi.org/10.1159/000335790. [DOI] [PubMed] [Google Scholar]
  57. Kraynov VS, Chamberlain C, Bokoch GM, Schwartz MA, Slabaugh S, Hahn KM. Localized Rac activation dynamics visualized in living cells. Science. 2000;290(5490):333–337. doi: 10.1126/science.290.5490.333. [DOI] [PubMed] [Google Scholar]
  58. Krugmann S, Jordens I, Gevaert K, Driessens M, Vandekerckhove J, Hall A. Cdc42 induces filopodia by promoting the formation of an IRSp53:Mena complex. Current Biology. 2001;11(21):1645–1655. doi: 10.1016/s0960-9822(01)00506-1. [DOI] [PubMed] [Google Scholar]
  59. Kurisu S, Takenawa T. The WASP and WAVE family proteins. Genome Biology. 2009;10(6):226. doi: 10.1186/gb-2009-10-6-226. http://dx.doi.org/10.1186/gb-2009-10-6-226. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Lang RA. Pathways regulating lens induction in the mouse. International Journal of Developmental Biology. 2004;48(8-9):783–791. doi: 10.1387/ijdb.041903rl. [DOI] [PubMed] [Google Scholar]
  61. Lang RA, Herman K, Reynolds AB, Hildebrand JD, Plageman TF., Jr p120-Catenin-dependent junctional recruitment of Shroom3 is required for apical constriction during lens pit morphogenesis. Development. 2014;141(16):3177–3187. doi: 10.1242/dev.107433. http://dx.doi.org/10.1242/dev.107433. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Lee C, Le MP, Wallingford JB. The shroom family proteins play broad roles in the morphogenesis of thickened epithelial sheets. Developmental Dynamics. 2009;238(6):1480–1491. doi: 10.1002/dvdy.21942. http://dx.doi.org/10.1002/dvdy.21942. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Lee-Hoeflich ST, Causing CG, Podkowa M, Zhao X, Wrana JL, Attisano L. Activation of LIMK1 by binding to the BMP receptor, BMPRII, regulates BMP-dependent dendritogenesis. EMBO Journal. 2004;23(24):4792–4801. doi: 10.1038/sj.emboj.7600418. http://dx.doi.org/10.1038/sj.emboj.7600418. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Leptin M. Gastrulation movements: The logic and the nuts and bolts. Developmental Cell. 2005;8(3):305–320. doi: 10.1016/j.devcel.2005.02.007. http://dx.doi.org/10.1016/j.devcel.2005.02.007. [DOI] [PubMed] [Google Scholar]
  65. Linseman DA, Loucks FA. Diverse roles of Rho family GTPases in neuronal development, survival, and death. Frontiers in Bioscience. 2008;13:657–676. doi: 10.2741/2710. [DOI] [PubMed] [Google Scholar]
  66. Liu H, Mohamed O, Dufort D, Wallace VA. Characterization of Wnt signaling components and activation of the Wnt canonical pathway in the murine retina. Developmental Dynamics. 2003;227(3):323–334. doi: 10.1002/dvdy.10315. http://dx.doi.org/10.1002/dvdy.10315. [DOI] [PubMed] [Google Scholar]
  67. Liu H, Thurig S, Mohamed O, Dufort D, Wallace VA. Mapping canonical Wnt signaling in the developing and adult retina. Investigative Ophthalmology & Visual Science. 2006;47(11):5088–5097. doi: 10.1167/iovs.06-0403. http://dx.doi.org/10.1167/iovs.06-0403. [DOI] [PubMed] [Google Scholar]
  68. Liu H, Xu S, Wang Y, Mazerolle C, Thurig S, Coles BL. Ciliary margin transdifferentiation from neural retina is controlled by canonical Wnt signaling. Developmental Biology. 2007;308(1):54–67. doi: 10.1016/j.ydbio.2007.04.052. http://dx.doi.org/10.1016/j.ydbio.2007.04.052. [DOI] [PubMed] [Google Scholar]
  69. Machacek M, Hodgson L, Welch C, Elliott H, Pertz O, Nalbant P, et al. Coordination of Rho GTPase activities during cell protrusion. Nature. 2009;461(7260):99–103. doi: 10.1038/nature08242. http://dx.doi.org/10.1038/nature08242. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Mallarino R, Grant PR, Grant BR, Herrel A, Kuo WP, Abzhanov A. Two developmental modules establish 3D beak-shape variation in Darwin’s finches. Proceedings of the National Academy of Sciences of the United States of America. 2011;108(10):4057–4062. doi: 10.1073/pnas.1011480108. http://dx.doi.org/10.1073/pnas.1011480108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Mann I. The development of the human eye. New York: Grune and Stratton; 1950. [Google Scholar]
  72. Martin AC, Kaschube M, Wieschaus EF. Pulsed contractions of an actinmyosin network drive apical constriction. Nature. 2009;457(7228):495–499. doi: 10.1038/nature07522. http://dx.doi.org/10.1038/nature07522. [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Martin-Blanco E, Pastor-Pareja JC, Garcia-Bellido A. JNK and decapentaplegic signaling control adhesiveness and cytoskeleton dynamics during thorax closure in Drosophila. Proceedings of the National Academy of Sciences of the United States of America. 2000;97(14):7888–7893. doi: 10.1073/pnas.97.14.7888. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. McAvoy JW. Cytoplasmic processes interconnect lens placode and optic vesicle during eye morphogenesis. Experimental Eye Research. 1980;31(5):527–534. doi: 10.1016/s0014-4835(80)80011-x. [DOI] [PubMed] [Google Scholar]
  75. McClay DR. The role of thin filopodiain motility and morphogenesis. Experimental Cell Research. 1999;253(2):296–301. doi: 10.1006/excr.1999.4723. http://dx.doi.org/10.1006/excr.1999.4723. [DOI] [PubMed] [Google Scholar]
  76. Mckeehan MS. Cytological aspects of embryonic lens induction in the chick. Journal of Experimental Zoology. 1951;117:31–64. [Google Scholar]
  77. Miki H, Sasaki T, Takai Y, Takenawa T. Induction of filopodium formation by a WASP-related actin-depolymerizing protein N-WASP. Nature. 1998;391(6662):93–96. doi: 10.1038/34208. [DOI] [PubMed] [Google Scholar]
  78. Miki H, Suetsugu S, Takenawa T. WAVE, a novel WASP-family protein involved in actin reorganization induced by Rac. The EMBO Journal. 1998;17(23):6932–6941. doi: 10.1093/emboj/17.23.6932. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Millard TH, Martin P. Dynamic analysis of filopodial interactions during the zippering phase of Drosophila dorsal closure. Development. 2008;135(4):621–626. doi: 10.1242/dev.014001. http://dx.doi.org/10.1242/dev.014001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  80. Moores SL, Sabry JH, Spudich JA. Myosin dynamics in live Dictyostelium cells. Proceedings of the National Academy of Sciences of the United States of America. 1996;93(1):443–446. doi: 10.1073/pnas.93.1.443. [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Nakano T, Ando S, Takata N, Kawada M, Muguruma K, Sekiguchi K, et al. Self-formation of optic cups and storable stratified neural retina from human ESCs. Cell Stem Cell. 2012;10(6):771–785. doi: 10.1016/j.stem.2012.05.009. http://dx.doi.org/10.1016/j.stem.2012.05.009. [DOI] [PubMed] [Google Scholar]
  82. Nimnual AS, Taylor LJ, Bar-Sagi D. Redox-dependent downregulation of Rho by Rac. Nature Cell Biology. 2003;5(3):236–241. doi: 10.1038/ncb938. http://dx.doi.org/10.1038/ncb938. [DOI] [PubMed] [Google Scholar]
  83. Nishimura T, Takeichi M. Shroom3-mediated recruitment of Rho kinases to the apical cell junctions regulates epithelial and neuroepithelial planar remodeling. Development. 2008;135(8):1493–1502. doi: 10.1242/dev.019646. [DOI] [PubMed] [Google Scholar]
  84. Nobes C, Hall A. Regulation and function of the Rho subfamily of small GTPases. Current Opinion in Genetics & Development. 1994;4(1):77–81. doi: 10.1016/0959-437x(94)90094-9. [DOI] [PubMed] [Google Scholar]
  85. Nobes CD, Hall A. Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell. 1995;81(1):53–62. doi: 10.1016/0092-8674(95)90370-4. [DOI] [PubMed] [Google Scholar]
  86. Nobes CD, Hall A. Rho GTPases control polarity, protrusion, and adhesion during cell movement. Journal of Cell Biology. 1999;144(6):1235–1244. doi: 10.1083/jcb.144.6.1235. [DOI] [PMC free article] [PubMed] [Google Scholar]
  87. Ohta Y, Hartwig JH, Stossel TP. FilGAP, a Rho- and ROCK-regulated GAP for Rac binds filamin A to control actin remodelling. Nature Cell Biology. 2006;8(8):803–814. doi: 10.1038/ncb1437. http://dx.doi.org/10.1038/ncb1437, ncb1437 [pii] [DOI] [PubMed] [Google Scholar]
  88. Pang SC, Daniels WH, Buck RC. Epidermal migration during the healing of suction blisters in rat skin: A scanning and transmission electron microscopic study. The American Journal of Anatomy. 1978;153(2):177–191. doi: 10.1002/aja.1001530202. http://dx.doi.org/10.1002/aja.1001530202. [DOI] [PubMed] [Google Scholar]
  89. Partridge MA, Marcantonio EE. Initiation of attachment and generation of mature focal adhesions by integrin-containing filopodia in cell spreading. Molecular Biology of the Cell. 2006;17(10):4237–4248. doi: 10.1091/mbc.E06-06-0496. http://dx.doi.org/10.1091/mbc.E06-06-0496. [DOI] [PMC free article] [PubMed] [Google Scholar]
  90. Passey S, Pellegrin S, Mellor H. What is in a filopodium? Starfish versus hedgehogs. Biochemical Society Transactions. 2004;32(Pt. 6):1115–1117. doi: 10.1042/BST0321115. http://dx.doi.org/10.1042/BST0321115. [DOI] [PubMed] [Google Scholar]
  91. Peng J, Wallar BJ, Flanders A, Swiatek PJ, Alberts AS. Disruption of the diaphanous-related formin Drf1 gene encoding mDia1 reveals a role for Drf3 as an effector for Cdc42. Current Biology. 2003;13(7):534–545. doi: 10.1016/s0960-9822(03)00170-2. [DOI] [PubMed] [Google Scholar]
  92. Pfister RR. The healing of corneal epithelial abrasions in the rabbit: A scanning electron microscope study. Investigative Ophthalmology. 1975;14(9):648–661. [PubMed] [Google Scholar]
  93. Plageman TF, Jr, Chauhan BK, Yang C, Jaudon F, Shang X, Zheng Y, et al. A Trio-RhoA-Shroom3 pathway is required for apical constriction and epithelial invagination. Development. 2011;138(23):5177–5188. doi: 10.1242/dev.067868. http://dx.doi.org/10.1242/dev.067868. [DOI] [PMC free article] [PubMed] [Google Scholar]
  94. Plageman TF, Jr, Chung MI, Lou M, Smith AN, Hildebrand JD, Wallingford JB, et al. Pax6-dependent Shroom3 expression regulates apical constriction during lens placode invagination. Development. 2010;137(3):405–415. doi: 10.1242/dev.045369. http://dx.doi.org/10.1242/dev.045369. [DOI] [PMC free article] [PubMed] [Google Scholar]
  95. Raftopoulou M, Hall A. Cell migration: Rho GTPases lead the way. Developmental Biology. 2004;265(1):23–32. doi: 10.1016/j.ydbio.2003.06.003. [pii] [DOI] [PubMed] [Google Scholar]
  96. Raich WB, Agbunag C, Hardin J. Rapid epithelial-sheet sealing in the Caenorhabditis elegans embryo requires cadherin-dependent filopodial priming. Current Biology. 1999;9(20):1139–1146. doi: 10.1016/S0960-9822(00)80015-9. http://dx.doi.org/10.1016/S0960-9822(00)80015-9. [DOI] [PubMed] [Google Scholar]
  97. Rajagopal R, Huang J, Dattilo LK, Kaartinen V, Mishina Y, Deng CX, et al. The type I BMP receptors, Bmpr1a and Acvr1, activate multiple signaling pathways to regulate lens formation. Developmental Biology. 2009;335(2):305–316. doi: 10.1016/j.ydbio.2009.08.027. http://dx.doi.org/10.1016/j.ydbio.2009.08.027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  98. Ridley AJ. Rho GTPases and actin dynamics in membrane protrusions and vesicle trafficking. Trends in Cell Biology. 2006;16(10):522–529. doi: 10.1016/j.tcb.2006.08.006. [DOI] [PubMed] [Google Scholar]
  99. Roffers-Agarwal J, Xanthos JB, Kragtorp KA, Miller JR. Enabled (Xena) regulates neural plate morphogenesis, apical constriction, and cellular adhesion required for neural tube closure in Xenopus. Developmental Biology. 2008;314(2):393–403. doi: 10.1016/j.ydbio.2007.12.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  100. Roh-Johnson M, Shemer G, Higgins CD, McClellan JH, Werts AD, Tulu US, et al. Triggering a cell shape change by exploiting preexisting actomyosin contractions. Science. 2012;335(6073):1232–1235. doi: 10.1126/science.1217869. http://dx.doi.org/10.1126/science.1217869. [DOI] [PMC free article] [PubMed] [Google Scholar]
  101. Rossman KL, Der CJ, Sondek J. GEF means go: Turning on RHO GTPases with guanine nucleotide-exchange factors. Nature Reviews Molecular Cell Biology. 2005;6(2):167–180. doi: 10.1038/nrm1587. http://dx.doi.org/10.1038/nrm1587. [DOI] [PubMed] [Google Scholar]
  102. Samakovlis C, Hacohen N, Manning G, Sutherland DC, Guillemin K, Krasnow MA. Development of the Drosophila tracheal system occurs by a series of morphologically distinct but genetically coupled branching events. Development. 1996;122(5):1395–1407. doi: 10.1242/dev.122.5.1395. [DOI] [PubMed] [Google Scholar]
  103. Sawyer JK, Harris NJ, Slep KC, Gaul U, Peifer M. The Drosophila afadin homologue Canoe regulates linkage of the actin cytoskeleton to adherens junctions during apical constriction. Journal of Cell Biology. 2009;186(1):57–73. doi: 10.1083/jcb.200904001. http://dx.doi.org/10.1083/jcb.200904001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  104. Sawyer JM, Harrell JR, Shemer G, Sullivan-Brown J, Roh-Johnson M, Goldstein B. Apical constriction: A cell shape change that can drive morphogenesis. Developmental Biology. 2010;341(1):5–19. doi: 10.1016/j.ydbio.2009.09.009. http://dx.doi.org/10.1016/j.ydbio.2009.09.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  105. Schmidt S, Diriong S, Mery J, Fabbrizio E, Debant A. Identification of the first Rho-GEF inhibitor, TRIPalpha, which targets the RhoA-specific GEF domain of Trio. FEBS Letters. 2002;523(1-3):35–42. doi: 10.1016/s0014-5793(02)02928-9. [DOI] [PubMed] [Google Scholar]
  106. Shaham O, Menuchin Y, Farhy C, Ashery-Padan R. Pax6: A multi-level regulator of ocular development. Progress in Retinal and Eye Research. 2012;31(5):351–376. doi: 10.1016/j.preteyeres.2012.04.002. http://dx.doi.org/10.1016/j.preteyeres.2012.04.002. [DOI] [PubMed] [Google Scholar]
  107. Smith AN, Miller LA, Radice G, Ashery-Padan R, Lang RA. Stage-dependent modes of Pax6-Sox2 epistasis regulate lens development and eye morphogenesis. Development. 2009;136(17):2977–2985. doi: 10.1242/dev.037341. http://dx.doi.org/10.1242/dev.037341. [DOI] [PMC free article] [PubMed] [Google Scholar]
  108. Snapper SB, Takeshima F, Anton I, Liu CH, Thomas SM, Nguyen D, et al. N-WASP deficiency reveals distinct pathways for cell surface projections and microbial actin-based motility. Nature Cell Biology. 2001;3(10):897–904. doi: 10.1038/ncb1001-897. [DOI] [PubMed] [Google Scholar]
  109. Sousa AD, Cheney RE. Myosin-X: A molecular motor at the cell’s fingertips. Trends in Cell Biology. 2005;15(10):533–539. doi: 10.1016/j.tcb.2005.08.006. http://dx.doi.org/10.1016/j.tcb.2005.08.006. [DOI] [PubMed] [Google Scholar]
  110. Steding G. Reasons for embryonic epithelial thickening. Acta Anatomica (Basel) 1967;68(1):37–67. [PubMed] [Google Scholar]
  111. Streit A. The cranial sensory nervous system: Specification of sensory progenitors and placodes. Cambridge, MA: StemBook; 2008. [PubMed] [Google Scholar]
  112. Takahashi K, Suzuki K. Membrane transport of WAVE2 and lamellipodia formation require Pak1 that mediates phosphorylation and recruitment of stathmin/Op18 to Pak1-WAVE2-kinesin complex. Cellular Signalling. 2009;21(5):695–703. doi: 10.1016/j.cellsig.2009.01.007. http://dx.doi.org/10.1016/j.cellsig.2009.01.007. [DOI] [PubMed] [Google Scholar]
  113. Tcherkezian J, Lamarche-Vane N. Current knowledge of the large RhoGAP family of proteins. Biology of the Cell. 2007;99(2):67–86. doi: 10.1042/BC20060086. http://dx.doi.org/10.1042/BC20060086. [DOI] [PubMed] [Google Scholar]
  114. Tilney LG, Gibbins JR. Microtubules and filaments in the filopodia of the secondary mesenchyme cells of Arbacia punctulata and Echinarachnius parma. Journal of Cell Science. 1969;5(1):195–210. doi: 10.1242/jcs.5.1.195. [DOI] [PubMed] [Google Scholar]
  115. Tokuo H, Ikebe M. Myosin X transports Mena/VASP to the tip of filopodia. Biochemical and Biophysical Research Communications. 2004;319(1):214–220. doi: 10.1016/j.bbrc.2004.04.167. http://dx.doi.org/10.1016/j.bbrc.2004.04.167. [DOI] [PubMed] [Google Scholar]
  116. Trimarchi JM, Cho SH, Cepko CL. Identification of genes expressed preferentially in the developing peripheral margin of the optic cup. Developmental Dynamics. 2009;238(9):2327–2329. doi: 10.1002/dvdy.21973. http://dx.doi.org/10.1002/dvdy.21973. [DOI] [PMC free article] [PubMed] [Google Scholar]
  117. Vasioukhin V, Fuchs E. Actin dynamics and cell-cell adhesion in epithelia. Current Opinion in Cell Biology. 2001;13(1):76–84. doi: 10.1016/s0955-0674(00)00177-0. [DOI] [PubMed] [Google Scholar]
  118. Villalonga P, Ridley AJ. Rho GTPases and cell cycle control. Growth Factors. 2006;24(3):159–164. doi: 10.1080/08977190600560651. [DOI] [PubMed] [Google Scholar]
  119. Wagner MC, Barylko B, Albanesi JP. Tissue distribution and subcellular localization of mammalian myosin I. Journal of Cell Biology. 1992;119(1):163–170. doi: 10.1083/jcb.119.1.163. [DOI] [PMC free article] [PubMed] [Google Scholar]
  120. Wakely J. Observations on the role of ectodermal spreading in the early stages of lens placode invagination in the chick embryo. Experimental Eye Research. 1984;38(6):627–636. doi: 10.1016/0014-4835(84)90181-7. [DOI] [PubMed] [Google Scholar]
  121. Wawersik S, Purcell P, Rauchman M, Dudley AT, Robertson EJ, Maas R. BMP7 acts in murine lens placode development. Developmental Biology. 1999;207(1):176–188. doi: 10.1006/dbio.1998.9153. [DOI] [PubMed] [Google Scholar]
  122. Westenskow P, Piccolo S, Fuhrmann S. Beta-catenin controls differentiation of the retinal pigment epithelium in the mouse optic cup by regulating Mitf and Otx2 expression. Development. 2009;136(15):2505–2510. doi: 10.1242/dev.032136. http://dx.doi.org/10.1242/dev.032136. [DOI] [PMC free article] [PubMed] [Google Scholar]
  123. Williams-Masson EM, Malik AN, Hardin J. An actin-mediated two-step mechanism is required for ventral enclosure of the C. elegans hypodermis. Development. 1997;124(15):2889–2901. doi: 10.1242/dev.124.15.2889. [DOI] [PubMed] [Google Scholar]
  124. Wolf C, Gerlach N, Schuh R. Drosophila tracheal system formation involves FGF-dependent cell extensions contacting bridge-cells. EMBO Reports. 2002;3(6):563–568. doi: 10.1093/embo-reports/kvf115. http://dx.doi.org/10.1093/embo-reports/kvf115. [DOI] [PMC free article] [PubMed] [Google Scholar]
  125. Wood W, Jacinto A, Grose R, Woolner S, Gale J, Wilson C, et al. Wound healing recapitulates morphogenesis in Drosophila embryos. Nature Cell Biology. 2002;4(11):907–912. doi: 10.1038/ncb875. [DOI] [PubMed] [Google Scholar]
  126. Xu J, Wang F, Van Keymeulen A, Herzmark P, Straight A, Kelly K, et al. Divergent signals and cytoskeletal assemblies regulate self-organizing polarity in neutrophils. Cell. 2003;114(2):201–214. doi: 10.1016/s0092-8674(03)00555-5. [DOI] [PubMed] [Google Scholar]
  127. Yeh TC, Ogawa W, Danielsen AG, Roth RA. Characterization and cloning of a 58/53-kDa substrate of the insulin receptor tyrosine kinase. Journal of Biological Chemistry. 1996;271(6):2921–2928. doi: 10.1074/jbc.271.6.2921. [DOI] [PubMed] [Google Scholar]
  128. Zhang H, Berg JS, Li Z, Wang Y, Lang P, Sousa AD, et al. Myosin-X provides a motor-based link between integrins and the cytoskeleton. Nature Cell Biology. 2004;6(6):523–531. doi: 10.1038/ncb1136. http://dx.doi.org/10.1038/ncb1136. [DOI] [PubMed] [Google Scholar]

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