Cell-cell adhesion in the cnidaria: insights into the evolution of tissue morphogenesis.
Adhesion between cells and the extracellular-matrix (ECM) surrounding them is vital for organismal morphogenesis, and thus the evolution of the structural components required for adhesion is intimately connected with the evolution of form. This review examines the evolution of the protein components responsible for cell adhesion, focusing on the recently sequenced genome of the cnidarian Nematostella vectensis Stephenson, 1935. Genomic approaches to identifying junctional components are providing important insights into the origins and evolution of the gene families involved in cell adhesion. Possession of complete genomic sequences provides unprecedented access to the complement of junctional components in taxa for which these resources are available.
Broadly speaking, there are two main categories of cellular morphologies in metazoans: epithelial and mesenchymal (Tyler, 2003). The differences between these grades of cellular organization are largely based on differences in their adhesive properties. An epithelium is defined as a sheet of polarized cells. The cells are joined by belt-like junctions around their apical margins, and ECM is typically present only apically and basally due to the close apposition of cells within the epithelium (though in acoel flatworms the basal lamina is absent; Rieger et al, 1991). Mesenchymal cells, in contrast, do not share the type of collective organization exhibited by epithelial cells. They are often isolated, with no direct neighbors. Rather than an organized belt of junctions restricted to particular membrane domains, they possess only spot-like junctions between the cell and the ECM, which can be present on any or all cell surfaces, or none. The conversion between epithelial and mesenchymal morphologies is involved in the development of many structures in the organism.
In addition to providing robust structural integrity to tissues, the organization of cells into epithelia partitions the organism into discrete compartments and thereby allows it to establish a specialized internal environment, which is required for the formation of complex organ systems. Cell-cell contacts are involved in this compartmentalization by preventing the diffusion of molecules across the epithelial layer, and are therefore important in the physiological function of the tissue. In sponges, epithelia form only transiently when required for a particular purpose, such as spicule secretion, and sponges are therefore not considered to be truly epithelial animals (Green and Bergquist, 1982). Cnidarians, on the other hand, are almost entirely epithelial, possessing both an outer epithelial layer and an inner epithelial digestive cavity protected from the external environment (Mergner, 1971). In bilaterians, interactions between epithelial and mesenchymal cells during development result in elaboration and further compartmentalization of the organism, which is required for proper organ formation, physiological function, and maintenance of homeostasis.
The conversion of cells from an epithelial to a mesenchymal morphology involves their detachment from the epithelium through an epithelial-to-mesenchymal transition (EMT), in which the ingressing cell dismantles its adhesive contacts, reorganizes its cytoskeleton to modify its shape, and migrates out of the plane of the epithelium (Shook and Keller, 2003). The mesenchymal cell can then migrate to its ultimate destination in the organism, where it differentiates or induces differentiation in neighboring cells (or both). EMT is utilized in a variety of developmental contexts, including gastrulation (Keller et al., 2003). Gastrulation, or the transition from a monolayered blastula to a multilayered gastrula, is the first morphogenetic event in embryogenesis, and results in the formation of the embryonic germ layers. Development and regulation of the various cell junctional complexes involved in cell adhesion, then, is of primary importance in the regulation of morphogenesis. Despite the diversity in morphology observed across the Metazoa, there are clear similarities in the types of cell junctions found in various taxa. Initially, these similarities were identified ul-trastructurally (e.g., Green and Flower, 1980; Green and Bergquist, 1982; Kachar et al., 1986; Fig. 1), but as the tools necessary to elucidate their molecular composition have become available, the morphological similarities have been found to reflect an underlying molecular similarity. This allows cell junctions to be categorized on the basis of their molecular and morphological structure.
[FIGURE 1 OMITTED]
Development and Regulation of Adhesive Contacts
The development of the various types of adhesive contacts is intimately tied to the development of cell polarity, as each of the junctional types discussed below are found in particular locations along the apico-basal axis of the epithelium (Fig. 2). A full discussion of the mechanisms underlying cell polarity and the genes involved in it is beyond the scope of this review, but some details are relevant for our discussion of adhesive complexes. In this section we describe the formation of the major junctional types and their regulation during development.
[FIGURE 2 OMITTED]
Early in development, cells of the cleavage-stage embryo undergo "epithelialization" and become adherent to one another. This early adhesion is largely based on transmembrane molecules of the cadherin superfamily, and it results in the formation of a monolayer embryo. Cadherins are calcium-dependent glycoproteins that mediate cell-cell interactions by binding to opposing cadherin molecules on neighboring cells. The cadherin superfamily is made up of the classical cadherins, desmosomal cadherins, protocadherins, and other cadherin-like proteins that do not fit into any of these subfamilies (Nollet et al., 2000; Fig. 3A). Because these categories were first identified in vertebrates, the classical cadherins are named on the basis of the tissues with which they are associated (e.g., E-cadherins in epithelia, N-cadherins in neural tissue). Adherens junctions (AJs) are defined by the presence of cadherins (Takeichi, 1991), and are the major junctional type required for cell-cell adhesion across the Metazoa (Oda et al., 2005). AJs can be organized either as focal spot junctions or as a belt-like zonula adherens (ZA) in mature epithelia. AJs are visible in transmission electron micrographs as narrow bands of electron-dense material flanking the plasma membranes of two adjacent cells (e.g., Spiegel and Howard, 1983; Fig. 1B).
[FIGURE 3 OMITTED]
Vertebrate classical cadherins consist of an N-terminal extracellular domain composed of five ectodomain (EC) modules (Takeichi, 1991). EC1 to EC4 are "cadherin repeats"--conserved domains that are common to all cadherin molecules, although the number of repeats differs across the cadherin subfamilies. EC5 in classical cadherins is known as the membrane proximal extracellular domain, and its sequence diverges somewhat from the other EC domains (Takeichi, 1991). Adjacent to EC5 is the transmembrane domain, followed by the C-terminal cytoplasmic domain that contains binding sites for cytoplasmic scaffold molecules of the catenin family. Vertebrate classical cadherins possess a conserved His-Ala-Val (HAV) sequence in their EC1 domain and are grouped together as "classical" or "type-I" cadherins on this basis (Blaschuk et al., 1990). Several other cadherins are very similar to the classical cadherins, but lack the HAV motif. These cadherins have been termed "atypical" or "type-II" cadherins (Tanihara et al., 1994). These family designations are also supported by phylogenetic analyses utilizing alignments of the EC1 domains from various cadherin molecules (Nollet et al., 2000).
Cadherins are linked to the cytoskeleton through the proteins of the catenin family: [alpha]-catenin, [beta]-catenin, and p120 catenin each contain a series of armadillo repeats, and all bind to the cytoplasmic domain of cadherin molecules (Hatzfeld, 1999; Steinberg and McNutt, 1999). [alpha] and [beta] catenin play structural roles in linking the cadherin complex to the actin cytoskeleton. The role of p120 catenin is less understood, but it probably involves regulating cadherin stability at the cell surface, as loss of cadherin-p120 binding results in endocytosis of E-cadherin in mammalian cells (Miyashita and Ozawa, 2007). Endocytosis of cadherins is a major way their subcellular localization is regulated (Ivanov et al., 2005), making modulation of this process crucial for the proper regulation of AJ complexes. Nonchordate cadherins have been identified (e.g., DE-cadherin in Drosophila) that possess the catenin-binding motif present in the cytoplasmic domain of chordate classical cadherins, and these have also been called "classical" cadherins on this basis, despite variations in extracellular domain structure (Oda et al., 2005).
Desmosomes constitute a second type of cadherin-based AJ present in vertebrates. They are related to typical AJs and in transmission electron micrographs exhibit a similar morphology, but they differ in the cytoplasmic scaffold molecules and cytoskeletal elements they are associated with. Their transmembrane molecules are the desmosomal cadherins: desmogleins and desmocollins (Nollet et al, 2000). Desmogleins and desmocollins are capable of binding to one another, and in fact there is evidence that heterodimers are preferred (Huber, 2003). Whereas the cadherins in most AJs are linked to the actin cytoskeleton through [alpha]-and [beta]-catenin, desmosomal cadherins recruit the armadillo proteins plakoglobin and plakophilin-1, which are vertebrate-specific gene duplications related to [beta]-catenin and are linked to the intermediate filament cytoskeleton either directly or via the linker protein desmoplakin (Schneider et al., 2003). Desmogleins and desmocollins possess the same 4-cadherin-repeat extracellular domain structure as type-I and type-II cadherins, though their cytoplasmic domains are unrelated. In addition, desmogleins exhibit longer cytoplasmic domains than desmocollins (Nollet et al., 2000).
Other cadherin-related proteins, including the protecadherins, Flamingo-like cadherins, and Fat-like cadherins, contain a variable number of cadherin repeats. For example, in vertebrates these cadherins possess between 6 and 34 repeats (Hong et al., 2004). Fat-like cadherins also contain a laminin A G repeat homology motif (LAG), along with EGF-like/cysteine-rich motifs (EGF) in their extracellular domain (Mahoney et al., 1991). Members of the Flamingo-like cadherin subfamily are distinguished by the presence of seven transmembrane domains, and they also contain EGF and LAG domains (Chae et al., 1999). Whether Fat- and Flamingo-like cadherins play a structural, adhesive role is not well understood, though they have been linked to signaling during tissue growth and planar cell polarity processes in both vertebrates and arthropods (Shimada et al., 2001; Castillejo-Lopez et al., 2004; Formstone and Mason, 2005; Willecke et al., 2006).
The early subcellular distribution of cadherin molecules is variable from taxon to taxon, but often more diffuse than it will ultimately be once the belt-like AJs of the ZA are formed. In the sea urchin embryo, for example, cadherin proteins are present throughout the cytoplasm and cell surface of cleavage-stage embryos, although they are enriched at sites of cell-cell contact (Miller and McClay, 1997). By the 16-cell stage, cadherin protein is restricted to lateral domains of cell-cell contact; by the blastula stage, it is further restricted to apical AJs. A similar pattern is seen in Caenorhabditis elegans embryos, with the cadherin HMR-1 initially present throughout the cytoplasm and plasma membrane of cleavage-stage blastomeres (but again enriched at sites of cell-cell contact), and restricted apically once AJs begin to form in the developing hypodermis (Costa et al., 1998). In Drosophila, the initiation of cellularization is accompanied by the development of AJs at the invaginating furrow canal (Tepass et al., 2001). These junctions move basally with the furrow canal as cellularization proceeds and are known as basal AJs. Apical spot AJs form around mid-cellularization, and are maintained in the apical third of the lateral cell membrane until gastrulation, when they form the belt-like AJs of the mature embryonic epithelium and the basal AJs are dismantled. In Nematostella vectensis, AJs are readily identified on transmission electron micrographs early in embryogenesis (Fritzenwanker et al., 2007). Spot AJs can be seen as early at the 16-cell stage; and by the blastula stage., belt AJs can be found at the apical cell margin. Smaller basolateral spot AJs are also present at this stage. Apical AJs persist through gastrulation, though are reduced in number in the invaginating endodermal cells (Kraus and Technau, 2006; Magie et al., 2007).
One of the molecules crucial for the proper apical positioning of AJs is the transmembrane protein Crumbs (Crb), which is localized to the apical membrane (Tepass, 1996). Crb consists of an extracellular domain with many cysteine-rich EGF-like repeats (30 in Drosophila). The cytoplasmic domain of Crb interacts with the membrane-associated guanylate kinase proteins (MAGUKs) encoded by the stardust (sdt) gene in Drosophila, along with another scaffold protein, Discs-lost (DIt), which contains four PDZ-domains (Medina et al., 2002). In Drosophila crb and sdt mutants, the apical spot AJs are not assembled into a functional ZA, but instead remain distributed as spot junctions in the lateral membrane (Gibson and Perrimon, 2003). Crb, Sdt, and Dlt homologs have been found in both C. elegans and mammalian genomes, indicating the degree to which this complex has been conserved (Bossinger et al., 2001).
Occluding junctions: septate and tight junctions
Septate and tight junctions (SJs and TJs, respectively) are included under one category because they are thought to serve analogous roles in the taxa they are found in, although their molecular composition and evolutionary origin may differ. Both TJs and SJs are thought to act as occluding junctions that restrict the paracellular diffusion of molecules across the epithelia in which they are found. They are quite different in morphology, however. TJs are located apical to AJs and consist of a series of discrete contact points between adjacent membranes where the intercellular space is completely obscured (Tsukita and Furuse, 1999; Fig. 1A). SJs, on the other hand, are located basal to AJs and are characterized by the presence of a series of ladder-like proteinaceous septa in the intercellular space (Green and Bergquist, 1982; Kachar et al., 1986; Fig. 1C). TJs are present exclusively in chordate taxa (with the possible exception of some arthropod nervous systems; Lane and Chandler, 1980), and are distinguished by the presence of transmembrane proteins of the occludin, claudin, and junction adhesion molecule (JAM) families (Gonzalez-Mariscal et al., 2003; Fig. 3C). Occludin and claudins constitute the primary TJ components, with JAM more restricted to cells of the immune system in mammals. Occludin and the claudins are "tetraspan" proteins, which refers to their four transmembrane domains (Furuse et al., 1993). Both N- and C-termini are oriented toward the cytoplasm, resulting in two extracellular loops. In Occludin, the loops are roughly the same size, whereas in claudins one loop is larger than the other. The C-terminus of Occludin is capable of interacting directly with actin filaments, PKC-, the tyrosine kinase Yes, and PI-3 kinase, as well as with the MAGUK proteins ZO-1, ZO-2, and ZO-3. Claudins, in contrast, do not bind actin directly but contain C-terminal PDZ (PSD-95/SAP90, Discs-large, ZO-1) motifs that bind the proteins ZO-1, ZO-2, and ZO-3. In mammals, the number of claudins that have been identified to date has reached 24 (Furuse and Tsukita, 2006).
As with AJs, TJ development is tied intimately to the development of cell polarity. During mouse development, TJs begin to form at the 8-cell stage (Fleming et al., 1989). The "epithelialization" of the embryo through expression of the cadherin/catenin complex results in the apico-basal polarization of the blastomeres. Formation of TJs then begins with the recruitment of ZO-1 to foci at the apicolateral cell contacts. By the 16-cell stage these ZO-1 containing foci expand into a belt-like morphology, and at the 32-cell stage Occludin and Claudin are added to the maturing TJ.
In another similarity with AJ development, the apical determinant Crb is also crucial for the development and localization of TJs in vertebrate epithelia, by recruiting the polarity proteins Par3, Par6, and associated proteins to the apical margin of the epithelium (Macara, 2004). PAR ("partition defective") proteins were first identified in C. elegans, where mutations in PAR genes result in a failure to properly specify the anterior-posterior axis in the early embryo (Kemphues et al., 1988). Par3 (known as Bazooka in Drosophila) and Par6 are PDZ-domain-containing proteins, and act as scaffolds (Etemad-Moghadam et al., 1995; Hung and Kemphues, 1999). Par6 forms a complex with the kinase aPKC, which has been shown to phosphorylate Par3, among other proteins (Lin et al., 2000). Recruitment of the Par3-Par6-aPKC complex is required for subsequent TJ morphogenesis.
SJs, so named because of the septa that characterize them in electron micrographs, are not found in chordate taxa, except between myelinating Schwann cells and the axons they encircle (termed paranodal junctions in this context; Hortsch and Margolis, 2003). Instead, they are found in many invertebrates and appear to serve the same occluding function as TJs in chordates. The molecular composition of SJs has been studied most extensively in Drosophila, and has led to the identification of a number of SJ components. It has been thought that TJs and SJs differ in their molecular composition, and in fact many Drosophila homologs of vertebrate TJ proteins such as PKC and ZO1-3 do not localize to the SJ, but instead to more apical membrane domains (Itoh et al., 1997; Harris and Peifer, 2005). Recently, however, claudins have been identified in Drosophila and C. elegans and found to be involved in the formation of SJs and paracellular barrier junctions, respectively, which suggests that TJs and SJs may be more closely related than previously thought (Asano et al., 2003; Wu et al., 2004). Alternatively, claudins may have been recruited independently to both junctional types. Another transmembrane protein found in SJs in Drosophila is Neurexin IV, a member of the Caspr (Cortactin-associated protein) family of neuronal cell surface proteins (Baumgartner et al., 1996; Fig. 3E). Caspr proteins have an extracellular domain containing a discoidin-like domain along with EGF and LAG domains. A Caspr homolog is also present in the paranodal SJs in vertebrates (Einheber et al., 1997). In Drosophila, additional SJ transmembrane proteins include cell adhesion molecules of the immunoglobulin (IgG) superfamily such as Fasciclin III and Neuroglian (Woods et al., 1997; Genova and Fehon, 2003; Faivre-Sarrailh et al., 2004). Proteins that interact with the cytoplasmic face of SJs include the MAGUK Discs-large (Dlg), members of the Protein 4.1/ Ezrin-moesin-radixin (ERM) family, and Scribble, a protein with multiple PDZ domains and leucine-rich repeats (Fehon et al., 1994; Bilder and Perrimon, 2000; Bossinger et al., 2001). ERM proteins are involved in linking the actin cytoskeleton to the junctional complex, whereas Dlg and Scrib play a more regulatory role and are required for the proper positioning of the SJ. Dlg is also generally involved in the establishment of epithelial polarity (Woods et al., 1996).
SJ development shares some important similarities with TJ development. As mentioned above, TJs are found in chordate taxa, but are absent from non-chordates. SJs, however, are present in some form across the Metazoa. SJs can form early in embryogenesis. During sea urchin development, for example, SJs form during the blastula stage basal to the apical AJs that also form at that stage (Spiegel and Howard, 1983). Morphologically, SJs form in a stepwise manner: components are assembled first into isolated plaques, which extend and link into the characteristic undulating pathways of the mature SJ (Lane and Swales, 1982). The number of parallel septae increases as the junction develops, until the SJ reaches its ultimate size. Despite the presence of SJs in Hydra, Nematostella does not appear to possess SJs early in embryogenesis (Magie et al., 2007). Given the presence of SJs in other anthozoans, however, it is likely that Nematostella does develop SJs by metamorphosis, though this has not been examined to date.
Gap junctions (GJs) are pore-containing cell-cell contacts that allow for the transport of small molecules from cell to cell and are important in cell-cell communication (Saez et al., 2003). The major pore-forming proteins in chordates are members of the connexin family (Sohl and Willecke, 2004). Connexins form hemichannels when six subunits oligomerize to form a hexameric ring, which initially adopts a closed configuration. Complete channels are then formed by the binding of two hemichannels, one from each of two adjacent cells (Fig. 1D). GJ formation thus requires prior cell-cell adhesion (Jongen et al., 1991). Connexins have not been found in invertebrates; instead, invertebrate GJs are composed of proteins initially known as innexins (invertebrate connexin; Panchin et al., 2000; Phelan and Starich, 2001). Following the identification of innexin homologs in vertebrates, however, proteins in this family were termed pannexins (Panchin, 2005). Pannexins, like connexins, possess four transmembrane domains and two extracellular loops, with both the N- and C-termini located in the cytoplasm, and they also form pores by organizing into hexameric rings (Fig. 3B). Pannexin homologs have been found in the genome of the hydrozoan cnidarian Hydra, though not in the anthozoan Nematostella vectensis (Alexopoulos et al., 2004). GJs have been observed ultrastructurally in Hydra and other hydrozoan cnidarians (e.g., Westfall et al., 1980). In the case of Nematostella (and anthozoans generally), GJs have also not been identified ultrastructurally (Mackie et al., 1984). There is, however, some evidence based on dye injection experiments that cells in the anthozoan Renilla koellikeri are coupled; and antibody recognition data suggest that a connexin-like gene exists in that organism, though clear GJs could not be found ultrastructurally (Germain and Anctil, 1996). If it were true that non-hydrozoan cnidarians lack GJs, the fact that GJs are abundant in hydrozoans as well as bilaterians would argue that junctions of this type have been secondarily lost in the other cnidarian taxa. At the minimum, GJs in anthozoans, if they exist, have been modified to such a degree as to be unrecognizable morphologically.
Proteins that interact with the cytoplasmic face of GJs include proteins from other types of junctions such as ZO-1 and members of the catenin family, as well as kinases such as Src, PKA, and PKC, and the microtubule components [alpha]-and [beta]-tubulin (Giepmans, 2004). Both connexins and pannexins interact with a similar complement of cytoplasmic proteins, highlighting the similarities between these junctional complexes despite the differences between connexins and pannexins at the amino acid level.
Focal adhesions (FAs) are spot junctions that mediate cell-ECM interactions, resulting in a link between the cytoskeleton of the cell and the ECM surrounding it (Fig. 1E). In a polarized epithelium, they are found at the basal margin of the epithelial cells and are involved in attachment of the epithelium to the basement membrane. In mesenchymal cells, they can be found on any cell surface that contacts ECM and are important in allowing the cell to migrate on that substrate. Their defining components are proteins of the integrin family (van der Flier and Sonnenberg, 2001). Integrins are transmembrane molecules that function as heterodimers between various [alpha] and [beta] forms (Fig. 3D). Integrins have been found in taxa from sponges to vertebrates, though the number of family members varies (Brower et al., 1997). Both the [alpha] and [beta] subunits are type-I transmembrane glycoproteins, with large extracellular domains, a single transmembrane domain, and short cytoplasmic domains. The [alpha] subunits have an extracellular N-terminal 7-bladed [beta]-propeller domain (Luo et al., 2007). Between the second and third "blades" of the propeller, many [alpha] subunits contain an "instead" (I) domain, also known as a von Willebrand factor A domain, which is the major site for ligand binding. The [beta] subunits also contain an I domain similar to that of the [alpha] I domain, except that it contains additional segments that are important for binding to the [alpha] subunit and in ligand-binding specificity. Integrin ligands include ECM components such as laminin, collagen, or fibronectin (Luo et al., 2007). A subset of these ligands contain an RGD (Arg-Gly-Asp) motif through which they interact with the integrin heterodimer (Luo et al., 2007), leading to the designation of this motif as a "disintegrin" domain.
Development of FAs occurs when cells come into contact with extracellular integrin ligands. Binding of integrins to their ligands induces clustering, either through multiple binding sites on the ligands themselves, or through conformational changes in the integrins that induce binding of crosslinking cytoplasmic components (Maheshwari et al., 2000). Once integrins have aggregated, they recruit cytoplasmic proteins such as paxillin and talin, which in turn recruit the organizing kinase Focal adhesion kinase (FAK; Mitra et al., 2005). FAK then phosphorylates the actin-binding proteins [alpha]-actinin and vinculin, resulting in the linkage of the actin cytoskeleton to the developing FA. FAs are involved in regulating cell shape and contractility by serving as anchor points between the cell and the surrounding ECM, and they thereby play important roles in cell motility and morphogenesis. The expression of integrins and other FA components has not yet been investigated in Nematostella, though integrins in other cnidarians are co-expressed with other FA components, suggesting that they may share a common molecular architecture (Reber-Muller et al. 2001). In support of this idea, Nematostella does possess homologs of at least some FA cytoplasmic components, including FAK, vinculin, and paxillin.
Regulation of adhesive contacts
Regulation of adhesive contacts after their initial formation is a critical aspect of the use of adhesion in morphogenesis. EMT is required for the development of mesenchyme, and the complementary process, the mesenchymal-to-epithelial transition (MET), is also used in the formation of some organs (e.g., Takahashi et al., 2005). Dismantling of the apical junctional complex during EMT is necessary for epithelial cells to detach from one another and migrate away as individuals (Boyer et al., 2000). In mammalian systems, both AJs and TJs disappear during EMT. This is accomplished, at least in part, by repressing the transcription of their respective adhesive proteins by transcription factors such as the DNA-binding zinc-finger protein Snail (Cano et al., 2000; Carver et al., 2001). Downregulation of components of the apical junctional complex through endocytosis and subsequent targeting of the protein complexes for degradation also contributes to this process, allowing for more precise temporal control of EMT (Ivanov et al., 2005).
EMT occurs at multiple times during development, including during gastrulation in many species. As the first morphogenetic process in the embryo, gastrulation is one of the central events in metazoan development. Strategies for transitioning from a monolayered blastula to a multilayered gastrula involve many types of cellular behaviors. These behaviors include invagination (the coordinated movement of sheets of cells into the interior of the embryo); epiboly (the spreading of cells over the surface of the embryo); delamination (mitoses in which the spindle is oriented perpendicular to the embryo surface, resulting in one daughter remaining on the surface and the other entering the blasto-coel); and ingression (during which cells detach from the epithelial layer of the blastula and migrate into the embryo as individuals; Keller et al., 2003). The ingression of cells into the blastocoel at gastrulation requires those cells to undergo EMT, making the regulation of cell adhesion crucial for the normal development of the organism.
In vertebrate embryos, the development of the somites and the formation of the neural crest are additional contexts in which the regulation of adhesion is critical for proper morphogenesis. In the case of the neural crest, the presumptive neural crest cells must undergo EMT to migrate out from the dorsal neural tube to reach their final positions in the embryo (Duband et al., 1995). Development of the somites involves both the formation of adhesive contacts and their breakdown. During somitogenesis in the early embryo, the newly specified somites begin as a mass of mesenchymal cells (Gossler and Hrabe de Angelis, 1998). Expression of ECM proteins and N-cadherin results in the reorganization of the outer cells of each somite into an epithelium (an MET; Linask et al., 1998). As the somites mature later in development, their ventral-medial cells (the sclerotome) undergo EMT, lose their epithelial contacts, and revert to a mesenchymal state. These mesenchymal cells will go on to form the cartilage of the vertebrae and ribs.
Once a cell undergoes EMT and becomes mesenchymal, it must be able to crawl upon the ECM surrounding it. Linking the cytoskeleton of the cell with the external ECM is the role of the integrin-based FAs. The ability of mesenchymal cells to migrate involves not only the assembly of FAs at the leading edge of the cell, but also their dismantling at the rear (Lauffenburger and Horwitz, 1996). In fact, repeatedly assembling FAs at one end of the cell and removing them at the other, coupled with actin polymerization in the direction of movement, is what drives the movement of the cell. Because of this, migrating mesenchymal cells can be described as "polarized" after a fashion, though it is not a stable polarization of the type seen in epithelial cells.
Evolution of Cell Adhesion
The evolution of cell adhesion is intimately connected with the evolution of multicellularity (Cereijido et al., 2004). The ability for cells to adhere to one another is of obvious importance in the development of multicellular forms, and in this sense the epithelium can be viewed as the fundamental metazoan innovation. The early evolution of the Metazoa, then, is essentially the evolution of the ability of cells to organize into epithelia, something that requires the cell-cell contacts discussed above. Examination of early-branching taxa such as cnidarians can provide insight into the nature of cell adhesion in these organisms, and thereby provide insight into the evolution of multicellularity itself. Further, the availability of whole genomes allows for the comparison of suites of genes involved in a particular process, which can provide a comprehensive view of the evolution of those gene networks.
During development in Nematostella, the first junctions that are observed in the embryo at the late cleavage stages are spot adherens junctions (Fritzenwanker et al., 2007). By the blastula stage, these spot junctions are elaborated into a belt-like zonula adherens. This pattern of junction development parallels in some respects the evolutionary history of cell-cell junctions, with poriferans exhibiting transient spot junctions, and belt-like junctions forming in cnidarian, ctenophore, and bilaterian taxa (Green and Bergquist, 1982; see Fig. 4). This observation is suggestive that perhaps the increases in anatomical complexity associated with bilaterian taxa were made possible by increases in the complexity of cell-cell junctional complexes.
[FIGURE 4 OMITTED]
When investigating the evolution of a particular process, it is helpful to examine an outgroup to the group under study, in order to polarize the direction of evolutionary change. In the case of the Metazoa, one such outgroup that has been examined is the choanoflagellates, which are single-celled protists hypothesized to be closely related to metazoan taxa. Expressed-sequence-tag sequences from the choanoflagellate species Monosiga brevicollis and Proterospongia-like sp. ATCC50818 reveal that, despite their unicellular nature, both these species contain cadherin genes (King et al., 2003). The cadherins identified are most closely related to the Flamingo and protocadherin subfamilies rather than to the "classical" cadherins. This suggests that the characteristic 5-EC domain structure of chordate classical cadherins is a modification specific to chordates, a hypothesis supported by the examination of the Drosophila melanogaster and Caenorhabditis elegans genomes, which has revealed that the cadherins in these species are more variable in their structure (Hill et al., 2001). A subset of cadherins identified in Drosophila and C. elegans do, however, possess a catenin binding motif in their cytoplasmic domain that is similar to that present in chordate classical cadherins, and because of this have also been termed "classical" cadherins (Oda and Tsukita, 1999).
Investigation of the cell adhesion genes in cnidarians has revealed that they possess many of the same proteins present in bilaterians. Searches of the Nematostella vectensis genome in particular reveal orthologs of most of the transmembrane adhesive proteins described above, with the notable exception of connexins/pannexins, claudins, and occludin. Nematostella also has a number of cadherin genes, which, like the choanoflagellate genes, are most similar to the Flamingo, Fat, and protocadherin subfamilies (Fig. 5). Chordate-like classical cadherins have not been found. On the basis of its domain structure, however, Nematostella [beta]-catenin is predicted to function in cell-cell adhesion as well as in transcriptional regulation, and would likely bind to a cadherin molecule to do so (Schneider et al., 2003). This raises the possibility that the ancestral cadherin gene was a Flamingo/Fat/protocadherin-like gene capable of mediating cell-cell adhesion and interacting with catenins. In support of this possibility, a BLAST search utilizing the catenin-binding sequence from the cytoplasmic domain of a chordate classical cadherin returns a hit against at least one of the Nematostella cadherin sequences (Nv_Cad4), albeit with a rather high E value (0.07). This sequence does not group with the classical cadherins, but is instead a Fat-like cadherin.
[FIGURE 5 OMITTED]
The apparent absence of connexins from non-chordate taxa suggests that they are an innovation of the Chordata. Pannexins, then, would be the ancestral transmembrane gap junction (GJ) protein. The relationship between connexins and pannexins is unclear, though they are not thought to share a common ancestor (Panchin, 2005). Perhaps the structural requirements of a pore-forming complex necessitate proteins with the same general domain organization shared by both connexins and pannexins, namely four trans membrane domains and two extracellular loops. Further studies documenting GJ genes in more taxa will be helpful in clarifying the relationship between these families.
The lack of Connexin and Pannexin genes in Nematostella is not surprising, given that GJs are thought to be absent in anthozoan cnidarians. The fact that Hydra does possess a pannexin gene, along with the ubiquitous presence of these genes in bilaterian taxa, argues that anthozoans have secondarily lost this junction type. Reports of dye coupling between cells in an anthozoan (Germain and Anctil, 1996), however, suggest that they may possess some type of GJ that diverged enough to be unrecognizable morphologically or molecularly. Alternatively, anthozoan cni darians may have evolved a new type of coupling junction that is evolutionary unrelated to GJs in other organisms. The significance of antibody-recognition data suggesting that the anthozoan Renilla koellikeri possesses a connexin-like gene is unclear. The identity of the antigen should be confirmed (e.g., through sequence analysis) before any conclusions are drawn. Further investigation of the molecular details of cnidarian cell biology will be necessary to clarify this issue.
Septate junctions (SJs) are widespread across the Bilateria and have been observed in various cnidarians, including the hydrozoan Hydra as well as anthozoans. The apparent lack of SJs in Nematostella embryos may indicate that Nematostella is simplified in this respect relative to these taxa, though a more detailed examination of Nematostella tissues (particularly polyps) for the presence of SJs is necessary. Additionally, on the basis of morphological criteria, the hydrozoan and bilaterian SJs are more similar to one another than either is to anthozoan SJs, arguing that the anthozoan SJs are derived structures and that the hydrozoan and bilaterian SJs may descend from a common ancestor (Green and Bergquist, 1982). Further studies detailing the molecular composition of the Hydra SJs compared with the bilaterian SJs will help determine whether this is actually the case. To date, a Hydra ortholog of ZO-1, a tight junction (TJ) and adherens junction (AJ) component in other organisms, has been isolated and localized to the apical margin of epithelial cells in the Hydra polyp (Fei et al., 2000). This is not where it would be predicted to be if it were involved in SJ function, though its interaction partners have not yet been identified. The availability of the Hydra genome, and one hopes, other cnidarian species in the future, will go a long way toward identifying commonalities and differences in intercellular junctional composition across the Cnidaria. Ctenophores do not have SJs, but instead possess a morphologically unique junctional structure located apical to AJs, similar to the location of TJs in vertebrates (Hernandez-Nicole, 1991). To date, no TJ components have been identified in ctenophores, although a complete genome has not been sequenced yet. It will be interesting to determine the molecular composition of these junctions.
Integrins have been found across the Metazoa, from sponges to vertebrates, reflecting how central the ability of a cell to interact with the extracellular matrix is to metazoan morphogenesis. Integrins exist as heterodimers, consisting of an [alpha] and a [beta] subunit, though the number of family members in different taxa varies. In humans, 18 [alpha] and 8 [beta] subunits combine to form 24 heterodimeric complexes (Luo et al., 2007). Drosophila melanogaster possesses 5 [alpha] and 1 [beta] resulting in five heterodimeric complexes, and in C. elegans 2 [alpha] and 1 [beta] are used in two heterodimers (Hughes, 2001). One integrin [alpha] and one [beta] have been cloned from the sponge Geodia cydonium, and a [beta] subunit has been cloned from another sponge, Ophlitaspongia tenuis (Brower et al., 1997; Wimmer et al., 1999). In the Cnidaria, [alpha] and [beta] subunits have been found in the hydrozoan Podocoryne carnea (Reber-Muller et al., 2001), and [beta] integrin subunits have also been identified in the coral Acropora millepora (Brower et al., 1997). Nematostella has at least two [alpha] and 5 [beta] integrin subunits, although it is not clear how many heterodimers are formed. The far greater number of integrin sequences in vertebrates indicates that the gene family has radiated in the chordates, as is the case for many vertebrate gene families. A phylogenetic analysis of integrin sequences indicates that the sponge and cnidarian [beta]-integrin sequences group into one clade with other [beta]-integrins as sister groups, suggesting a cnidarian-specific radiation of [beta]-integrin genes (Fig. 6A). This is consistent with [beta]-integrin genes in other taxa, which also group into lineage-specific clades (Ewan et al., 2005). The sponge and cnidarian [alpha]-integrin sequences do not show this pattern, but instead group as sister to the PS2 group (Pc_[alpha], Nv_[alpha].b, Gc_[alpha]) and within the PS1 group (Nv_[alpha].a), suggesting that these types of [alpha]-integrins radiated after the split of the cnidarians from the last common metazoan ancestor (Fig. 6B). An examination of the [alpha]-integrin sequences failed to reveal the presence of an I-domain, consistent with the hypothesis that this domain is a chordate-specific innovation (Ewan et al., 2005).
In any examination of the ways cell adhesion genes have evolved within the Metazoa, the fundamental question is their origin. The perhaps surprising result that choanoflagellates, which are unicellular protists, possess cadherin genes indicates that adhesion genes were present in the ancestor to the Metazoa (King et al., 2003). The question then becomes, what was their function in a unicellular context? Future work in choanoflagellates will help clarify this issue, though we can speculate that these genes played a role similar to their function in metazoans. Perhaps cell adhesion genes in unicellular organisms evolved to stabilize interactions between mating pairs, or to permit colonial association of individuals, or to facilitate prey capture by allowing the cell to adhere to its prey. Alternatively, perhaps they played a role in signaling, as both cadherins and integrins have been shown to do in modern animals, and were secondarily recruited to junctional complexes. In each case, proteins that evolved for one purpose in the unicellular organism were evolutionarily co-opted for another purpose in order to construct a multicellular organism.
The genomic approach holds promise for clarifying evolutionary relationships among genes in early-branching taxa. It will be interesting to compare, for example, the recently completed genomes of sponges and placozoans with the cnidarians and ctenophores, and ultimately with bilaterians. The more genomes that are sequenced, the more resolution we will have as to how these various gene families have changed over evolutionary time, allowing for the evolution of the incredibly varied forms seen across the Metazoa.
CRAIG R. MAGIE AND MARK Q. MARTINDALE *
Kewalo Marine Laboratory, Pacific Biomedical Research Center, University of Hawai'i Honolulu, Hawaii 96813
Received 15 November 2007; accepted 6 February 2008.
* To whom correspondence should be addressed. E-mail: email@example.com
Abbreviations: AJ, adherens junction; EC, ectodomain; ECM, extracellular matrix; EMT, epithelial-to-mesenchymal transition; FA, focal adhesion; GJ, gap junction; MET, mesenchymal-to-epithelial transition; SJ, septate junction; TJ, tight junction; ZA, zonula adherens.
The authors thank members of the Martindale lab for helpful discussions, and two anonymous reviewers for constructive suggestions on the manuscript.
Alexopoulos, H., A. Bottger, S. Fischer, A. Levin, A. Wolf, T. Fujisawa, S. Hayakawa, T. Gojobori, J. A. Davies, C. N. David, and J. P. Bacon. 2004. Evolution of gap junctions: the missing link? Curr. Biol. 14: R879-880.
Asano, A., K. Asano, H. Sasaki, M. Furuse, and S. Tsukita. 2003. Claudins in Caenorhabditis elegans: their distribution and barrier function in the epithelium. Curr. Biol. 13: 1042-1046.
Baumgartner, S., J. T. Littleton, K. Broadie, M. A. Bhat, R. Harbecke, J. A. Lengyel, R. Chiquet-Ehrismann, A. Prokop, and H. J. Bellen. 1996. A Drosophila neurexin is required for septate junction and blood-nerve barrier formation and function. Cell 87: 1059-1068.
Bilder, D., and N. Perrimon. 2000. Localization of apical epithelial determinants by the basolateral PDZ protein Scribble. Nature 403: 676-680.
Blaschuk, O. W., R. Sullivan, S. David, and Y. Pouliot. 1990. Identification of a cadherin cell adhesion recognition sequence. Dev. Biol. 139: 227-229.
Bossinger, O., A. Klebes, C. Segbert, C. Theres, and E. Knust. 2001. Zonula adherens formation in Caenorhabditis elegans requires dlg-1, the homologue of the Drosophila gene discs large. Dev. Biol. 230: 29-42.
Boyer, B., A. M. Valles, and N. Edme. 2000. Induction and regulation of epithelial-mesenchymal transitions. Biochem. Pharmacol. 60: 1091-1099.
Brower, D. L., S. M. Brower, D. C. Hayward, and E. E. Ball. 1997. Molecular evolution of integrins: genes encoding integrin beta subunits from a coral and a sponge. Proc. Natl. Acad. Sci. USA 94: 9182-9187.
Cano, A., M. A. Perez-Moreno, I. Rodrigo, A. Locascio, M. J. Blanco, M. G. del Barrio, F. Portillo, and M. A. Nieto. 2000. The transcription factor snail controls epithelial-mesenchymal transitions by repressing E-cadherin expression. Nat. Cell Biol. 2: 76-83.
Carver, E. A., R. Jiang, Y. Lan, K. F. Oram, and T. Gridley. 2001. The mouse snail gene encodes a key regulator of the epithelial-mesenchymal transition. Mol. Cell. Biol. 21: 8184-8188.
Castillejo-Lopez, C., W. M. Arias, and S. Baumgartner. 2004. The fat-like gene of Drosophila is the true orthologue of vertebrate fat cadherins and is involved in the formation of tubular organs. J. Biol. Chem. 279: 24034-24043.
Cereijido, M., R. G. Contreras, and L. Shoshani. 2004. Cell adhesion, polarity, and epithelia in the dawn of metazoans. Physiol. Rev. 84: 1229-1262.
Chae, J., M. J. Kim, J. H. Goo, S. Collier, D. Gubb, J. Charlton, P. N. Adler, and W. J. Park. 1999. The Drosophila tissue polarity gene starry night encodes a member of the protocadherin family. Development 126: 5421-5429.
Costa, M., W. Raich, C. Agbunag, B. Leung, J. Hardin, and J. R. Priess. 1998. A putative catenin-cadherin system mediates morphogenesis of the Caenorhabditis elegans embryo. J. Cell Biol. 141: 297-308.
Duband, J. L., F. Monier, M. Delannet, and D. Newgreen. 1995. Epithelium-mesenchyme transition during neural crest development. Acta Anat. (Basel) 154: 63-78.
Einheber, S., G. Zanazzi, W. Ching, S. Scherer, T. A. Milner, E. Peles, and J. L. Salzer. 1997. The axonal membrane protein Caspr, a homologue of neurexin IV, is a component of the septate-like paranodal junctions that assemble during myelination. J. Cell Biol. 139: 1495-1506.
Etemad-Moghadam, B., S. Guo, and K. J. Kemphues. 1995. Asymmetrically distributed PAR-3 protein contributes to cell polarity and spindle alignment in early C. elegans embryos. Cell 83: 743-752.
Ewan, R., J. Huxley-Jones, A. P. Mould, M. J. Humphries, D. L. Robertson, and R. P. Boot-Handford. 2005. The integrins of the urochordate Ciona intestinalis provide novel insights into the molecular evolution of the vertebrate integrin family. BMC Evol. Biol. 5: 31.
Faivre-Sarrailh, C., S. Banerjee, J. Li, M. Hortsch, M. Laval, and M. A. Bhat. 2004. Drosophila contactin, a homolog of vertebrate contactin, is required for septate junction organization and paracellular barrier function. Development 131: 4931-4942.
Fehon, R. G., I. A. Dawson, and S. Artavanis-Tsakonas. 1994. A Drosophila homologue of membrane-skeleton protein 4.1 is associated with septate junctions and is encoded by the coracle gene. Development 120: 545-557.
Fei, K., L. Yan, J. Zhang, and M. P. Sarras, Jr. 2000. Molecular and biological characterization of a zonula occludens-1 homologue in Hydra vulgaris, named HZO-1. Dev. Genes Evol. 210: 611-616.
Fleming, T. P., J. McConnell, M. H. Johnson, and B. R. Stevenson. 1989. Development of tight junctions de novo in the mouse early embryo: control of assembly of the tight junction-specific protein. ZO-1. J. Cell Biol. 108: 1407-1418.
Formstone, C. J., and I. Mason. 2005. Combinatorial activity of Flamingo proteins directs convergence and extension within the early zebrafish embryo via the planar cell polarity pathway. Dev. Biol. 282: 320-335.
Fritzenwanker, J. H., G. Genikhovich, Y. Kraus, and U. Technau. 2007. Early development and axis specification in the sea anemone Nematostella vectensis. Dev. Biol. 310: 264-279.
Furuse, M., and S. Tsukita. 2006. Claudins in occluding junctions of humans and flies. Trends Cell Biol. 16: 181-188.
Furuse, M., T. Hirase, M. Itoh, A. Nagafuchi, S. Yonemura, and S. Tsukita. 1993. Occludin: a novel integral membrane protein localizing at tight junctions. J. Cell Biol. 123: 1777-1788.
Genova, J. L., and R. G. Fehon. 2003. Neuroglian, Gliotactin, and the [Na.sup.+]/[K.sup.+] ATPase are essential for septate junction function in Drosophila. J. Cell Biol. 161: 979-989.
Germain, G., and M. Anctil. 1996. Evidence for intercellular coupling and connexin-like protein in the luminescent endoderm of Renilla koellikeri (Cnidaria, Anthozoa). Biol. Bull. 191: 353-366.
Gibson, M. C., and N. Perrimon. 2003. Apicobasal polarization: epithelial form and function. Curr. Opin. Cell Biol. 15: 747-752.
Giepmans, B. N. 2004. Gap junctions and connexin-interacting proteins. Cardiovasc. Res. 62: 233-245.
Gonzalez-Mariscal, L., A. Betanzos, P. Nava, and B. E. Jaramillo. 2003. Tight junction proteins. Prog. Biophys. Mol. Biol. 81: 1-44.
Gossler, A., and M. Hrabe de Angelis. 1998. Somitogenesis. Curr. Top. Dev. Biol. 38: 225-287.
Green, C. R., and P. R. Bergquist. 1982. Phylogenetic relationships within the Invertebrata in relation to the structure of septate junctions and the development of 'occluding' junctional types. J. Cell Sci. 53: 279-305.
Green, C. R., and N. E. Flower. 1980. Two new septate junctions in the phylum Coelenterata. J. Cell Sci. 42: 43-59.
Harris, T. J., and M. Peifer. 2005. The positioning and segregation of apical cues during epithelial polarity establishment in Drosophila. J. Cell Biol. 170: 813-823.
Hatzfeld, M. 1999. The armadillo family of structural proteins. Int. Rev. Cytol. 186: 179-224.
Hernandez-Nicole, M. L. 1991. Ctenophora. Pp. 359-418 in Microscopic Anatomy of Invertebrates, Vol. 2, Placozoa, Porifera, Cnidaria, and Ctenophora, F. W. Harrison and J. A. Westfall, eds. Wiley-Liss, New York.
Hill, E., I. D. Broadbent, C. Chothia, and J. Pettitt. 2001. Cadherin superfamily proteins in Caenorhabditis elegans and Drosophila melanogaster. J. Mol. Biol. 305: 1011-1024.
Hong, J. C., N. V. Ivanov, P. Hodor, M. Xia, N. Wei, R. Blevins, D. Gerhold, M. Borodovsky, and Y. Liu. 2004. Identification of new human cadherin genes using a combination of protein motif search and gene finding methods. J. Mol. Biol. 337: 307-317.
Hortsch, M., and B. Margolis. 2003. Septate and paranodal junctions: kissing cousins. Trends Cell Biol. 13: 557-561.
Huber, O. 2003. Structure and function of desmosomal proteins and their role in development and disease. Cell Mol. Life Sci. 60: 1872-1890.
Huelsenbeck, J. P., and F. Ronquist. 2001. MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics 17: 754-755.
Hughes, A. L., 2001. Evolution of the integrin alpha and beta protein families. J. Mol. Evol. 52: 63-72.
Hung, T. J., and K. J. Kemphues. 1999. PAR-6 is a conserved PDZ domain-containing protein that colocalizes with PAR-3 in Caenorhabditis elegans embryos. Development 126: 127-135.
Itoh, M., A. Nagafuchi, S. Moroi, and S. Tsukita. 1997. Involvement of ZO-1 in cadherin-based cell adhesion through its direct binding to alpha catenin and actin filaments. J. Cell Biol. 138: 181-192.
Ivanov, A. I., A. Nusrat, and C. A. Parkos. 2005. Endocytosis of the apical junctional complex: mechanisms and possible roles in regulation of epithelial barriers. Bioessays 27: 356-365.
Jongen, W. M., D. J. Fitzgerald, M. Asamoto, C. Piccoli, T. J. Slaga, D. Gros, M. Takeichi, and H. Yamasaki. 1991. Regulation of connexin 43-mediated gap junctional intercellular communication by [Ca.sup.2+] in mouse epidermal cells is controlled by E-cadherin. J. Cell Biol. 114: 545-555.
Kachar, B., N. A. Christakis, T. S. Reese, and N. J. Lane. 1986. The intramembrane structure of septate junctions based on direct freezing. J. Cell Sci. 80: 13-28.
Keller, R., L. A. Davidson, and D. R. Shook. 2003. How we are shaped: the biomechanics of gastrulation. Differentiation 71: 171-205.
Kemphues, K. J., J. R. Priess, D. G. Morton, and N. S. Cheng. 1988. Identification of genes required for cytoplasmic localization in early C. elegans embryos. Cell 52: 311-320.
King, N., C T. Hittinger, and S. B. Carroll. 2003. Evolution of key cell signaling and adhesion protein families predates animal origins. Science 301: 361-363.
Kraus, Y., and U. Technau. 2006. Gastrulation in the sea anemone Nematostella vectensis occurs by invagination and immigration: an ultrastructural study. Dev. Genes Evol. 216: 119-132.
Lane, N. J., and H. J. Chandler. 1980. Definitive evidence for the existence of tight junctions in invertebrates. J. Cell Biol. 86: 765-774.
Lane, N. J., and L. S. Swales. 1982. Stages in the assembly of pleated and smooth septate junctions in developing insect embryos. J. Cell Sci. 56: 245-262.
Lauffenburger, D. A., and A. F. Horwitz. 1996. Cell migration: a physically integrated molecular process. Cell 84: 359-369.
Lin, D., A. S. Edwards, J. P. Fawcett, G. Mbamalu, J. D. Scott, and T. Pawson. 2000. A mammalian PAR-3-PAR-6 complex implicated in Cdc42/RAC 1 and aPKC signalling and cell polarity. Nat. Cell Biol. 2: 540-547.
Linask, K. K., C. Ludwig, M. D. Han, X. Liu, G. L. Radice, and K. A. Knudsen. 1998. N-cadherin/catenin-mediated morphoregulation of somite formation. Dev. Biol. 202: 85-102.
Luo, B. H., C. V. Carman, and T. A. Springer. 2007. Structural basis of integrin regulation and signaling. Annu. Rev. Immunol. 25: 619-647.
Macara, I.G. 2004. Parsing the polarity code. Nat. Rev. Mol. Cell Biol. 5: 220-231.
Mackie, G. O., P. A. V. Anderson, and C. L. Singla. 1984. Apparent absence of gap junctions in two classes of Cnidaria. Biol. Bull. 167: 120-123.
Magie, C. R., M. Daly, and M. Q. Martindale. 2007. Gastrulation in the cnidarian Nematostella vectensis occurs via invagination not ingression. Dev. Biol. 305: 483-497.
Maheshwari, G., G. Brown, D. A. Lauffenburger, A. Wells, and L. G. Griffith. 2000. Cell adhesion and motility depend on nanoscale RGD clustering. J. Cell Sci. 113 Pt 10: 1677-1686.
Mahoney, P. A., U. Weber, P. Onofrechuk, H. Biessmann, P. J. Bryant, and C. S. Goodman. 1991. The fat tumor suppressor gene in Drosophila encodes a novel member of the cadherin gene superfamily. Cell 67: 853-868.
Medina, E., C. Lemmers, L. Lane-Guermonprez, and A. Le Bivic. 2002. Role of the Crumbs complex in the regulation of junction formation in Drosophila and mammalian epithelial cells. Biol. Cell 94: 305-313.
Mergner, H. 1971. Cnidaria. Pp. 1-84 in Experimental Embryology of Marine and Fresh-water Invertebrates, G. Reverberi, ed. North-Holland Publishing, London.
Miller, J. R., and D. R. McClay. 1997. Changes in the pattern of adherens junction-associated beta-catenin accompany morphogenesis in the sea urchin embryo. Dev. Biol. 192: 310-322.
Mitra, S. K., D. A. Hanson, and D. D. Schlaepfer. 2005. Focal adhesion kinase: in command and control of cell motility. Nat. Rev. Mol. Cell. Biol. 6: 56-68.
Miyashita, Y., and M. Ozawa. 2007. Increased internalization of p120-uncoupled E-cadherin and a requirement for a dileucine motif in the cytoplasmic domain for endocytosis of the protein. J. Biol. Chem. 282: 11540-11548.
Nollet, F., P. Kools, and F. van Roy. 2000. Phylogenetic analysis of the cadherin superfamily allows identification of six major subfamilies besides several solitary members. J. Mol. Biol. 299: 551-572.
Oda, H., and S. Tsukita. 1999. Nonchordate classic cadherins have a structurally and functionally unique domain that is absent from chordate classic cadherins. Dev. Biol. 216: 406-422.
Oda, H., K. Tagawa, and Y. Akiyama-Oda. 2005. Diversification of epithelial adherens junctions with independent reductive changes in cadherin form: identification of potential molecular synapomorphies among bilaterians. Evol. Dev. 7: 376-389.
Panchin, Y. V. 2005. Evolution of gap junction proteins--the pannexin alternative. J. Exp. Biol. 208: 1415-1419.
Panchin, Y., I. Kelmanson, M. Matz, K. Lukyanov, N. Usman, and S. Lukyanov. 2000. A ubiquitous family of putative gap junction molecules. Curr. Biol. 10: R473-474.
Phelan, P., and T. A. Starich. 2001. Innexins get into the gap. Bioes-says 23: 388-396.
Reber-Muller, S., R. Studer, P. Muller, N. Yanze, and V. Schmid. 2001. Integrin and talin in the jellyfish Podocoryne carnea. Cell Biol. Int. 25: 753-769.
Rieger, R. M., S. Tyler, J. P. S. Smith III, and G. E. Rieger. 1991. Platyhelminthes: Turbellaria. Pp. 7-140 in Microscopic Anatomy of Invertebrates, Vol. 3, Platyhelminthes and Nemertinea, F. W. Harrison and B. J. Bogitsh, eds. Wiley-Liss, New York.
Saez, J. C., V. M. Berthoud, M. C. Branes, A. D. Martinez, and E. C. Beyer. 2003. Plasma membrane channels formed by connexins: their regulation and functions. Physiol. Rev. 83: 1359-1400.
Schneider, S. Q., J. R. Finnerty, and M. Q. Martindale. 2003. Protein evolution: structure-function relationships of the oncogene beta-catenin in the evolution of multicellular animals. .J. Exp. Zool. B Mol. Dev. Evol. 295: 25-44.
Shimada, Y., T. Usui, S. Yanagawa, M. Takeichi, and T. Uemura. 2001. Asymmetric colocalization of Flamingo, a seven-pass transmembrane cadherin, and Dishevelled in planar cell polarization. Curr. Biol. 11: 859-863.
Shook, D., and R. Keller. 2003. Mechanisms, mechanics and function of epithelial-mesenchymal transitions in early development. Mech. Dev. 120: 1351-1383.
Sohl, G., and K. Willecke. 2004. Gap junctions and the connexin protein family. Cardiovasc. Res. 62: 228-232.
Spiegel, E., and L. Howard. 1983. Development of cell junctions in sea-urchin embryos. J. Cell Sci. 62: 27-48.
Steinberg, M. S., and P. M. McNutt. 1999. Cadherins and their connections: adhesion junctions have broader functions. Curr. Opin. Cell Biol. 11: 554-560.
Swofford, D. L. 2002. PAUP*. Phylogenetic Analysis Using Parsimony (*and Other Methods). Version 4. Sinauer Associates, Sunderland, MA.
Takahashi, Y., Y. Sato, R. Suetsugu, and Y. Nakaya. 2005. Mesenchymal-to-epithelial transition during somitic segmentation: a novel approach to studying the roles of Rho family GTPases in morphogenesis. Cells Tissues Organs 179: 36-42.
Takeichi, M. 1991. Cadherin cell adhesion receptors as a morphogenetic regulator. Science 251: 1451-1455.
Tanihara, H., K. Sano, R. L. Heimark, T. St John, and S. Suzuki. 1994. Cloning of five human cadherins clarifies characteristic features of cadherin extracellular domain and provides further evidence for two structurally different types of cadherin. Cell Adhes. Commun. 2: 15-26.
Tepass, U. 1996. Crumbs, a component of the apical membrane, is required for zonula adherens formation in primary epithelia of Drosophila. Dev. Biol. 177: 217-225.
Tepass, U., G. Tanentzapf, R. Ward, and R. Fehon. 2001. Epithelial cell polarity and cell junctions in Drosophila Annu. Rev. Genet. 35: 747-784.
Tsukita, S., and M. Furuse. 1999. Occludin and claudins in tight-junction strands: leading or supporting players? Trends Cell Biol. 9: 268-273.
Tyler, S. 2003. Epithelium--the primary building block for metazoan complexity. Integr. Comp. Biol. 43: 55-63.
van der Flier, A., and A. Sonnenberg. 2001. Function and interactions of integrins. Cell Tissue Res. 305: 285-298.
Westfall, J. A., J. C. Kinnamon, and D. E. Sims. 1980. Neuro-epitheliomuscular cell and neuro-neuronal gap junctions in Hydra. J. Neurocytol. 9: 725-732.
Whelan, S., and N. Goldman. 2001. A general empirical model of protein evolution derived from multiple protein families using a maximum-likelihood approach. Mol. Biol. Evol. 18: 691-699.
Willecke, M., F. Hamaratoglu, M. Kango-Singh, R. Udan, C. L. Chen, C. Tao, X. Zhang, and G. Halder. 2006. The fat cadherin acts through the hippo tumor-suppressor pathway to regulate tissue size. Curr. Biol. 16: 2090-2100.
Wimmer, W., B. Blumbach, B. Diehl-Seifert, C. Koziol, R. Batel, R. Steffen, I. M. Muller, and W. E. Muller. 1999. Increased expression of integrin and receptor tyrosine kinase genes during autograft fusion in the sponge Geodia cydonium. Cell Adhes. Commun. 7: 111-124.
Woods, D. F., C. Hough, D. Peel, G. Callaini, and P. J. Bryant. 1996. Dlg protein is required for junction structure, cell polarity, and proliferation control in Drosophila epithelia. J. Cell Biol. 134: 1469-1482.
Woods, D. F., J. W. Wu, and P. J. Bryant. 1997. Localization of proteins to the apico-lateral junctions of Drosophila epithelia. Dev. Genet. 20: 111-118.
Wu, V. M., J. Schulte, A. Hirschi, U. Tepass, and G. J. Beitel. 2004. Sinuous is a Drosophila claudin required for septate junction organization and epithelial tube size control. J. Cell Biol. 164: 313-323.
|Printer friendly Cite/link Email Feedback|
|Author:||Magie, Craig R.; Martindale, Mark Q.|
|Publication:||The Biological Bulletin|
|Date:||Jun 1, 2008|
|Previous Article:||Does the high gene density in the sponge NK homeobox gene cluster reflect limited regulatory capacity?|
|Next Article:||Genomic survey of candidate stress-response genes in the estuarine anemone Nematostella vectensis.|