Chapter 3 Acquisition of Membrane Polarity in Epithelial Tube Formation

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T H R E E

Acquisition of Membrane Polarity in Epithelial Tube Formation: Patterns, Signaling Pathways, Molecular Mechanisms, and Disease Fernando Martı´n-Belmonte and Alejo E. Rodrı´guez-Fraticelli Contents 1. Introduction 2. Architecture of Polarized Cells in Epithelial Organs 2.1. Differentiated plasma membrane domains define epithelial architecture 2.2. Cell–cell junctions and polarity complexes 3. Epithelial Morphogenesis into Tubes 3.1. Model systems for studying tubulogenesis 3.2. Formation of epithelial tubes follows diverse patterns 4. De Novo Formation of Tubes: Conserved Pathways and Molecular Mechanisms 4.1. Initial cell–cell and cell–ECM interactions drive the formation of AJs and the orientation of the axis of polarity 4.2. Vesicle trafficking, membrane separation, and lumen coalescence in lumen formation 4.3. The role of PtdIns in the definition of membrane identity 5. Epithelial Polarity and Disease 5.1. Cell polarity and cancer 5.2. Trafficking disorders 5.3. Cytoskeletal and phosphoinositide related disorders 5.4. Cystic diseases of the kidney 6. Concluding Remarks Acknowledgments References

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Centro de Biologı´a Molecular Severo Ochoa, Consejo Superior de Investigaciones Cientı´ficas-UAM, Madrid 28049, Spain International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02003-0

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2009 Elsevier Inc. All rights reserved.

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Abstract Epithelia coordinate the polarity of individual cells, in space and time, with surrounding cells and the extracellular matrix (ECM) to organize threedimensional structures that shape tissues and organs. One of the most important features of epithelial polarization is the asymmetric distribution of membrane surfaces with the apical surface facing a lumen or outside of the organism, and a basolateral surface facing other cells and ECM. This chapter discuss the processes required for the acquisition of the asymmetric distribution of membrane surfaces during morphogenesis, which include trafficking pathways, vesiclesorting machineries, formation of junctional and polarity complexes, and the establishment of signaling networks. In addition, different mechanisms and patterns are described for forming luminal spaces, and how alterations in cell polarity are associated with important human diseases such as cancer.

1. Introduction Many of our most important organs display a complex tissue organization made of a highly intricate network of branched tubes, which are required for the transport and distribution of liquids and nutrients throughout the body. Tubular organs develop following different strategies, which generate a remarkable structural and cellular diversity: different tube sizes, shapes, and connecting patterns (Hogan and Kolodziej, 2002; Lubarsky and Krasnow, 2003). These tubes are mainly composed of monolayers of highly polarized epithelial cells surrounding a central lumen. One of the most important features of epithelial cells is the existence of different plasma membrane domains, apical and basolateral, characterized by their distinct composition of lipids and proteins. Multiple cellular processes are required for establishing apical–basal polarity, including polarized vesicular transport, polarization of the cytoskeleton, and the proper establishment of cell adhesion and cell junction complexes. At specific stages of embryonic development and in some cancers, epithelial cells can undergo an epithelial-to-mesenchymal transition (EMT) that causes the loss of cell–cell adhesions and polarity markers, which results in the activation of a migratory phenotype. Therefore, the integrity of epithelial polarity plays an essential role in tumor progression and health (Debnath and Brugge, 2005; Yamada and Cukierman, 2007). Furthermore, many human diseases such as polycystic kidney disease, atherosclerotic heart disease, and faciogenital dysplasia derive from defects in epithelial tubular organization. A molecular understanding of tube morphogenesis could lead to new ways of diagnosing and treating these conditions. In this review paper, we focus on the acquisition of membrane polarity during epithelial tube formation. In particular, we will review different patterns that epithelial organs follow during morphogenesis, as well as the

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recent discoveries in the signaling pathways and the molecular mechanisms of these processes. Additionally, we will review how alterations in the process of acquisition of membrane polarity during morphogenesis are associated with important human diseases such as cancer.

2. Architecture of Polarized Cells in Epithelial Organs 2.1. Differentiated plasma membrane domains define epithelial architecture The epithelial plasma membrane is divided into three surfaces: the apical membrane, the free surface facing the lumen; the basal membrane, which is in contact with the tissues underneath; and the lateral surfaces that connect neighboring cells through a set of specialized cellular junctions that anchor cells to one another. These cellular junctions provide barrier function, and preserve the integrity and the different composition of the membranes by blocking the movement of proteins and outer-leaflet lipids (Mostov et al., 2003). The apical membrane is often covered by abundant microvilli, and appears to be the more specialized domain, since it contains most of the proteins required for organ-specific functions, such as terminal digestion and nutrient absorption or resorption. Generally, the apical plasma membrane is enriched in PtdIns-4,5-p2, sphingomyelin-containing glycolipids, cholesterol, and glycolipid-anchored proteins. By contrast, the basolateral membrane is enriched in PtdIns 3,4,5 p3 and carries most of the constitutive functions of the cells (e.g., cholesterol uptake, growth factor receptor, etc.). To maintain cell polarity and play their specific functions, epithelial cells have to ensure proper delivery of apical and basolateral cargos to their respective target location. The sorting of membrane proteins into distinct transport carriers occurs, either in the trans-Golgi network (TGN) during direct biosynthetic delivery, or in the common recycling endosomes (CRE) during indirect biosynthetic delivery or cargo-recycling to the plasma membrane (Folsch, 2008; Rodriguez-Boulan et al., 2005). Polarized protein delivery is regulated by sorting signals contained within the proteins themselves, which are recognized by specific sorting machineries. Some of the components of these specific sorting machineries have been identified in recent years, providing crucial information about the molecular mechanism that govern the protein traffic in epithelial tissues (Folsch, 2008; Mostov et al., 2003; Rodriguez-Boulan et al., 2005). Figure 3.1 summarizes the different trafficking routes and sorting mechanisms in epithelial cells that include: biosynthetic, endocytic, recycling, and transcytotic. Biosynthetic routes provide newly synthesized proteins to the apical and basolateral membranes. After the synthesis of the proteins, they are transported along

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Lumen Apical membrane

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Figure 3.1 Trafficking pathways in polarized epithelial cells. Biosynthetic pathways allow the transport of newly synthesized proteins to the apical and basolateral surfaces from the TGN. The biosynthetic pathway to the basolateral surface could be direct or indirect through the common recycling endosome (CRE). See Folsch (2008) for details. Apical and basolateral endocytosis allows the internalization of surface proteins to the early endosomes; the apical early endosome (AEE) or the basolateral early endosome (BEE). The proteins at the early endosomes can be: (A) recycled back to the surface using the recycling pathways at the apical, through the apical recycling endosome (ARE), and/or basolateral surfaces the CRE; (B) send to the opposite surface using the pathway for transcytosis from the basolateral to the apical surface; (C) or send to the late endosomes (LE) for degradation.

the secretory pathway (ER, Golgi, and TGN), and sorted into carriers to the different domains at either the TGN, or the endosomes. Proteins at the plasma membrane are rapidly endocytosed and delivered to the early endosomes where they follow the endocytic route or, after passing through the CRE, are recycled back to the cell surface (recycling), or transported across the cell to the opposite plasma membrane (transcytosis). The importance of these pathways varies with the type of epithelial cell, but they must be finely regulated in order to induce and maintain the steady-state polarity of the cells. The transport of proteins along these trafficking routes is regulated by sorting signals present in the proteins themselves and recognized by specific sorting machineries.

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2.1.1. Sorting signals and sorting machineries It was initially proposed that whereas the basolateral sorting was dependent on cytoplasmic signals, the apical sorting was a ‘‘default’’ pathway. This hypothesis originated in experiments in which the mutation of the basolateral signals (described below) induced the missorting of the proteins to the apical domain (Matter and Mellman, 1994). However, further work has proved the existence of apical sorting information that typically consists of ectodomain, membrane, or cytoplasmic signals. The first apical signal characterized was the glycosylphosphatidylinositol (GPI) anchors present in certain proteins (GPI-anchored proteins) (Lisanti et al., 1988). The evidence for these results came from a series of experiments in which the recombinant addition of the GPI anchor to secretory proteins resulted in the apical localization of the chimeric proteins (Brown et al., 1989; Lisanti et al., 1989). A second group of apical sorting signals includes N-linked or O-linked glycans, present in the exoplasmic region of many glycoproteins. A third group of apical sorting signals is encoded by the protein sequences themselves (Rodriguez-Boulan et al., 2005). One example is the transmembrane domain of the influenza virus protein hemaglutinin (HA), a prototypical apical targeted protein, which contains apical sorting information (Tall et al., 2003). Finally, it has been recently described that apical sorting information can also be encoded by cytoplasmic and exoplasmic protein domains present in the apical proteins (Marzolo et al., 2003; Takeda et al., 2003). A common requirement for apical sorting seems to be a clustering of the apical proteins into specific membrane domains, perhaps with the help of lectins that recognize N- or O-linked glycans, for direct delivery from the TGN to the apical membrane (Delacour et al., 2005; Fiedler and Simons, 1995) or, due to the ability of some apical directed proteins such as GPI-anchored proteins, to oligomerize during their passage through the Golgi complex (Paladino et al., 2004). Additionally, these apical clustering could be mediated by either lipid-raft domains or nonraft carriers. The lipidraft hypothesis (van Meer and Simons, 1988) postulates that apical targeted proteins are clustered and incorporated in transport vesicles due to their affinity for microdomains enriched in glycosphigolipids and cholesterol. Different proteins have been postulated to promote the clustering of lipidrafts, including the MAL/VIP17 (Alonso and Millan, 2001) and the caveolin family of proteins (Simons and Ikonen, 1997), which induce the formation of large amounts of cytoplasmic membrane vesicle structures when overexpressed in insect cells (Li et al., 1996; Puertollano et al., 1997). The association of the MAL/VIP17 family of proteins with lipid-rafts and their biological function seems to be related to its MARVEL domain, which is also present in physins, gyrins, and occludin families (Sanchez-Pulido et al., 2002). The function of the MARVEL domain could be related to cholesterol-rich membrane apposition events in a variety of cellular

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processes, such as biogenesis of vesicular transport carriers or tight junction regulation. The role of MAL/VIP17 and other members of the family in raftassociated vesicle transport has been extensively addressed in epithelial cells (Cheong et al., 1999; de Marco et al., 2002; Martin-Belmonte et al., 2000, 2001, 2003; Puertollano et al., 1999). However, the specific role for caveolin protein in raft-associated vesicle traffic still needs to be clarified (Manninen et al., 2005; Scheiffele et al., 1998). Some apically transported proteins, such as gp114, the neurotrophin receptor p75, and lactase-phorizin hydrolase (LPH), are included in nonraft vesicle carriers (Delacour et al., 2006). These nonraft vesicles, however, need also to be clustered for apical delivery. In fact, recent data has shown that Galectin-3 can form oligomers, and is involved in the clustering and apical delivery of nonraft associated glycoproteins to the apical surface and to membrane polarization of mouse enterocytes in vivo (Delacour et al., 2006, 2007, 2008). Interestingly, it appears that nonraft and raft dependent carriers require the presence of adaptor proteins at the TGN for apical delivery. The phosphatidylinositol(4)phosphate (PI(4)P) binding protein FAPP2 has been recently described to be required for apical delivery of the raft-associated protein HA and GPI-anchored proteins, and also for nonraft-associated vesicles (Vieira et al., 2005, 2006). FAPP2 seems to be needed for the transport of glycosylceramide from the cis-Golgi to the TGN, where it is incorporated into glycosphingolipids (D’Angelo et al., 2007). The authors speculate with the possibility that FAPP2 coordinates glycosphingolipid synthesis with apical transport. The basolateral sorting information, by contrast, is composed of sorting peptides included in the cytoplasmic tails of the basolateral targeted transmembrane proteins. Basolateral sorting signals, first described for the polymeric IgA receptor (pIgR) (Casanova et al., 1991; Mostov et al., 1986), are typically tyrosine-based (YXX) or leucine-based (mono- or di-leucine) peptide motifs (Rodriguez-Boulan et al., 2005). They are usually dominant over apical sorting signals, which explain the fact that when they are removed, certain basolateral proteins are missorted to the apical surface (Folsch, 2008). These peptidic sorting signals are recognized by cytosolic adaptor proteins, which in general form heterotetramers and interact with the vesicle coating protein clathrin, also required for basolateral distribution (Deborde et al., 2008). There are four types of complexes AP-1, AP-2, AP-3, and AP-4, and five types of adaptin proteins. Each complex consist of two large subunits (a, g, E, d, and b, 1
4), a medium subunit (m, 1–4), and a small subunit (s, 1–4). The tyrosine sorting signal (YXX) is recognized by the m subunit, whereas the di-L motif is recognized by the g/s1 or d/s3 subunits ( Janvier et al., 2003). The interaction of the adaptor proteins with the cargo and the clathrin coats induces the clustering of the basolateral proteins into clathrin-coated pits, which are subsequently budded into the cytoplasm for basolateral distribution. However, the current knowledge

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about the specificity of the adaptor proteins in basolateral trafficking is still very poor. Furthermore, the only AP complex that has been proven to be associated with basolateral sorting identified so far is the AP–1B complex. The AP–1B complex is composed of the m1b subunit, that determines exclusively the polarity of tyrosine-based signals, and it is only expressed in a number of polarized epithelial cells (Folsch et al., 1999; Gan et al., 2002). The current knowledge about the transcytosis signals is very limited, except for pIgR, which has been extensively studied for many years (Mostov et al., 2003). Recent findings suggest the possibility that the inactivation of basolateral sorting signals, when the proteins have already reached the basolateral surface could induce the activation of apical delivery signals, and thus, mediate the transcytosis of proteins that present both sorting information from the basolateral-to-the-apical surface (Anderson et al., 2005). Furthermore, this potential mechanism for transcytosis would be defined by the dominant hierarchy of basolateral signals over the apical signal described before, which would allow the initial delivery of the proteins to the basolateral domain. 2.1.2. Membrane fusion machinery The tethering (or docking) and fusion of transport vesicles with apical or basolateral domains are essential during exocytosis, and are needed for the acquisition of cell polarity and membrane identity (Wu et al., 2008). However, little is known about the mechanisms that control all these events during exocytosis. The exocyst protein complex function has been reported to be involved in the tethering, docking, and fusion of post-Golgi vesicles with the plasma membrane in polarized cells (Fig. 3.2). The most recent information from several model systems has demonstrated that the three families of small GTPases (Rab, Ral, and Rho) regulate the function of the exocyst complex, which is composed of eight subunits conserved from yeast to mammalian cells: Sec3, Sec5, Sec6, Sec8, Sec10, Sec15, Exo70, and Exo84 (Wu et al., 2008). The initial vesicle-docking event is regulated by Rab and Ral GTPases, perhaps by promoting exocyst assembly. There is evidence that the exocyst assembly is regulated by Ral and this function, like that of Rab GTPases, is first required for vesicle tethering rather than fusion (Moskalenko et al., 2002, 2003). Assembly is followed by local activation of the exocyst complex by active, GTP-bound, Cdc42 or TC10 GTPases. Exocyst activation results in the stimulation of a downstream fusion activity, probably by promoting assembly of active t-SNARE heterodimers. The presence of active t-SNARE proteins results in the fusion of the secretory vesicles at the site of exocyst activation. In epithelial cells, the exocyst is localized in the Golgi apparatus, the TGN, RE, and the junctional complex, and is proposed to promote the targeting and fusion of biosynthetic and endocytic recycling cargo carriers with the basolateral plasma membrane domain, possibly at sites near the tight junction (Folsch et al., 2003;

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Figure 3.2 Model for basolateral vesicle docking and fusion regulated by small GTPases and the exocyst. See Wu et al. (2008) for more information. The initial vesicle-docking or tethering event is regulated by Rab8 and RalA GTPases by promoting exocyst assembly. The association of the exocyst subunits with the vesicle or plasma membrane in this diagram is speculative. This is followed by local activation of the exocyst complex by GTP-bound Cdc42 Rho GTPase. Exocyst activation results in a stimulation of downstream fusion activity, probably by promoting assembly of active t-SNAREs heterodimers (SNAP23 and Syntaxin 4). The presence of active t-SNARE dimers with ts-VAMP, the v-SNARE, results in SNARE-mediated fusion of the vesicles with the basolateral membrane at the site of exocyst activation.

Grindstaff et al., 1998; Yeaman et al., 2001, 2004). More recent results have shown that the exocyst could function in several endocytic pathways as well, including basolateral recycling, apical recycling, and basolateralto-apical transcytosis. The latter was selectively dependent on interactions between the small GTPase Rab11a and Sec15A (Oztan et al., 2007). In the apical membrane, current information about the machinery for vesicle fusion is limited. The annexins family of proteins, which associate with the plasma membrane in a Ca2þ and negative phospholipids dependent manner (Rescher and Gerke, 2004), could be part of the machinery that mediates the fusion of apically derived vesicles with this plasma membrane domain. In particular, annexin A2 forms heterotetramers at the plasma membrane with its ligand p11, and it is enriched at the apical domain through its association with PtdIns(4,5)p2 (Martin-Belmonte et al., 2007; Rescher et al., 2004). The delivery of sucrase-isomaltase to the apical domain in Madin–Darby Canine Kidney (MDCK) cells requires annexin A2 ( Jacob et al., 2004). Furthermore, recent results have shown that Annexin A2 is required for the formation of the apical membrane and the lumen in MDCK cysts in a pathway that also involved Cdc42 (Martin-Belmonte et al., 2007).

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As mentioned above, the last step of fusion of vesicles with a target plasma membrane is mediated by the SNARE complex, denominated v-SNAREs, and t-SNAREs on the vesicle and target membrane, respectively. In polarized epithelial cells, apical and basolateral vesicles contain different v-SNAREs (such as TI-VAMP for apical, VAMP8 and cellubrevin for basolateral transport).The t-SNAREs are localized to the apical (syntaxin-3 and SNAP23), and basolateral (syntaxin-4 and SNAP23) plasma membranes. Loss-of-function or mislocalization of SNAREs leads to a concomitant disruption of plasma membrane delivery of the apical or basolateral vesicle population (ter Beest et al., 2005).

2.2. Cell–cell junctions and polarity complexes Epithelial polarity in multicellular organisms is also regulated by the formation of cell–cell junctions between cells, as well as by the presence of polarity complexes conserved throughout evolution. The polarity program is partially executed in epithelial cells by the competition and sequestration of these polarity complexes in different subcellular domains. 2.2.1. Junctional complexes All along the lateral membrane that connects neighboring cells, the apical– basal axis is delimited by junctional complexes. In vertebrates, these complexes include apical tight junctions (TJs), laterally localized adherent junctions (AJs), desmosomes and the connect cells with the ECM. Analogous structures exist in the Drosophila epithelium with some small differences. The Drosophila equivalents of the TJ, known as the septate junction, are localized basal, rather than apical to the AJ, although their cellular functions appear to be similar. For the purpose of this review, we will concentrate our discussion on vertebrate TJs and AJs. TJs serve not only to establish an apical–basal barrier that inhibits the diffusion of solutes across the epithelial layer (gate function), but they also restrict the movement of proteins and outer-leaflet lipids between the apical and the basolateral membranes (fence function) (Matter and Balda, 2003). The TJs are composed primarily of the membrane-bound junctional adhesion molecules ( JAMs), claudins and occludins, which are connected to the cytoskeleton through the PDZ containing proteins zonula occludens 1–3 (ZO-1, ZO-2, and ZO-3). These adapters also recruit regulatory proteins, such as protein kinases, phosphatases, small GTPases, and transcription factors to the TJs. This protein scaffolding facilitates the assembly of highly ordered structures that regulate epithelial cell polarity, proliferation, and differentiation (Kohler and Zahraoui, 2005). AJs perform multiple functions including initiation and stabilization of cell–cell adhesion, regulation of the actin cytoskeleton, intracellular signaling, and transcriptional regulation. The central part of the AJs includes

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interactions among transmembrane glycoproteins of the classical cadherin superfamily, such as E-cadherin, and the catenin family members including p120-catenin, b-catenin, and a-catenin. Together, these proteins control the formation, maintenance, and function of AJs (Hartsock and Nelson, 2008). E-cadherin is the major transmembrane protein of the epithelial AJs, and initiates intercellular contacts through trans-pairing between cadherins on opposing cells (Gumbiner, 2005). Classical cadherins also bind directly and indirectly to many cytoplasmic proteins, particularly members of the catenin family, which locally regulate the organization of the actin cytoskeleton, cadherin stability, and intracellular signaling pathways that control gene transcription (Perez-Moreno and Fuchs, 2006). Formation of the AJs leads to assembly of the TJs, but E-cadherin is not required to maintain TJs organization (Capaldo and Macara, 2007). Recent results have shown that an hepatic cell line unable to form AJs can form functional TJs in a delayed manner (Theard et al., 2007). 2.2.2. Polarity complexes Throughout evolution and the development of highly complex multicellular organisms, the requirement for cell asymmetry has remained essential. Given this, it is not surprising that many of the protein networks that have evolved to regulate cell polarity in Caenorhabditis elegans, Drosophila melanogaster are conserved through all eukaryotic species. In fact, each protein present in the polarity complexes involved in regulating cell polarity in C. elegans and Drosophila has highly conserved mammalian homologues. Over the past 15 years, the identification and analysis of either single or multiple mammalian homologues of the Drosophila polarity proteins have served to validate a conserved functional role for these proteins in regulating epithelial cell polarity. Furthermore, the manipulation in cultured mammalian cells has allowed us to gain new insights into the physical and functional interactions of the polarity proteins, and the coordinated development of apical– basal polarity in epithelium that was not possible in other in vivo models. Par4, also called LKB1 in mammalians, is mutated in Peutz–Jeghers syndrome (PJS), causing a predisposition to benign and malignant epithelial tumors (discussed in Chapter 5 of this volume). Activation of LKB1 by STRAD in mammalian intestinal epithelial cells in culture has demonstrated an essential role in cell polarity: cells with activated Par4 become polarized in the absence of cell–cell and cell–extracellular-matrix interactions, form an actin-rich apical brush border, and localize the TJs component ZO-1, and the AJs component p120 around it (Baas et al., 2004). LKB1 phosphorylates many kinases, including both the polarity protein, Par1 (Fig. 3.3) and AMPK. One particularly interesting finding is the involvement of LKB1 and AMP Kinase (AMPK) as master regulators of the energy state of the cell. During energy stress, LKB1 and AMPK are vital for epithelial polarization in both Drosophila and cultured mammalian cells, but apparently, they are

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Figure 3.3 The signaling pathways regulated by the polarity complexes for epithelial polarity in mammalian cells. The acquisition of apical–basolateral polarity during epithelial morphogenesis requires the activity of several polarity complexes. Phosphorylation and binding of Par5 (black) provides a mechanism for mutual exclusion for different PAR proteins. The basolateral determinant Par1 (green) that has diffused onto the apical domain is phosphorylated by Par6/aPKC (white), which inhibits the Par1 kinase activity and induces binding to Par5, and release into the cytoplasm. The apical determinant Par3 (orange) that diffuses into the basolateral domain is phosphorylated by Par1, which induces binding of Par5 and release into the cytoplasm. STRAD (brown) phosphorylates LKB1 (light green), which in turn activates Par1. The basolateral determinant Lgl (light blue) that has diffused onto the apical domain is phosphorylated by aPKC and induces its release from the cell cortex into the cytoplasm, although the exact mechanism is still unkonwn. aPKC phosphorilates Crumbs (purple) at the apical cortex to control the extension of the apical surface and/or apical junction assembly, together with PatJ (grey) and Pals1 (light brown), forming the CRB complex. Lgl, Dlg (yellow) and scribble (light purple) form the SCRB complex to define the basolateral domain by inhibiting Par complex and the CRBs complex. The localization of Par6/aPKC to the apical cortex and TJ is mediated by activated Cdc42-GTP (blue) or Par3, respectively.

less necessary when the energy and ATP level of the cell is high (Lee et al., 2007; Mirouse et al., 2007; Zhang et al., 2006b; Zheng and Cantley, 2007). Par1 is a Ser/Thr kinase that is localized to the basolateral membrane and excludes apical determinants, such as Par3, from this domain (Fig. 3.3). In addition, Par1 determines the organization of microtubules in mammalian cells, which in turn establishes the position of the luminal surface (Cohen et al., 2004, 2007). High Par1 activity converts columnar epithelial cells with vertical microtubules and an apical luminal surface into a hepatic type of epithelial cells, with horizontal microtubules and lumens

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that form between adjacent cells. In addition, high Par1 activity also causes the direct transport of apical proteins to the apical domain be modified to transcytotic route (Cohen et al., 2004), and it might regulate the docking and fusion of vesicles at the plasma membrane, at least in yeast cells (Elbert et al., 2005). Apical–basolateral polarization is essentially mediated by three interacting protein complexes: (1) Par/aPKC complex, consisting of Par3 (Baz), Par6, and atypical protein kinase C (aPKC); (2) Crumbs complex, consisting of Crumbs, Stardust, PATJ and several recently identified components, such as Yurt and Tuberous sclerosis (TSC) (Laprise et al., 2006; Massey-Harroche et al., 2007); and (3) the Scribble complex, consisting of Scribble, Discs large (Dlg), and Lethal giant larvae (Lgl) (Fig. 3.3). These complexes have been reviewed in detail elsewhere (Goldstein and Macara, 2007; Wang and Margolis, 2007). The Par/aPKC complex is involved in polarity and spatial organization in almost all metazoan cells, whereas the Crumbs complex is more specific to epithelial cells. The functions of the Scrib complex in mammals include exocyst-dependent basolateral exocytosis, and apical junction regulation (mScrib inhibits junction formation at the basolateral domain) (Qin et al., 2005; Yamanaka and Ohno, 2008; Zhang et al., 2005). Even though the molecular nature of these relationships is mostly unknown, these three complexes interact by a system of mutual exclusion to define the apical and basolateral surfaces of epithelial cells in Drosophila (Bilder et al., 2003; Tanentzapf and Tepass, 2003), and they could function similarly in mammalian cells (Fig. 3.3). The Par/aPKC complex is a master regulator of polarity (Munro, 2006). Mammalian Par3 is localized to TJs through the interaction with JAM at the apical/lateral boundary (Izumi et al., 1998), and functions in their assembly (Chen and Macara, 2005), whereas Par6/aPKC maintains the integrity of the apical domain (Martin-Belmonte et al., 2007). Par6 acts as a targeting subunit for aPKC, and it recruits both Crumbs complex (Hurd et al., 2003b; Lemmers et al., 2004) and Lgl (Scribble complex) as substrates (Betschinger et al., 2005). CRB controls the extension of the apical membrane (Macara, 2004), whereas the Par3-mediated phosphorilation of Lgl restricts the localization of Lgl to the basolateral domain. On the other hand, the Scribble complex suppresses apical membrane identity in the basolateral domain by inhibiting the Par3 complex (Tanentzapf and Tepass, 2003). How these opposing activities lead to the coalescence of AJs into the mature zonula adherens is not known, although it could involve regulation of the polarized transport of membrane proteins. Lgl homologues interact with Myosin II (a regulator of the actin cytoskeleton), with components of the exocyst complex (which is involved in transporting proteins to the plasma membrane), and with the fragile X-associated protein FMR1 (which is involved in transport and translational control of specific mRNAs). Finally, recent data has implicated PKA in TJ assembly (Kohler and Zahraoui, 2005). PKA activity is required to recruit claudin1 and ZO1, and

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inhibits the association of vasodilator-stimulated protein (VASP), indicating that PKA signaling is important for the assembly of functional TJs. In addition, cAMP and PKA activity stimulate apically directed transcytosis and secretion in epithelial cells, and budding of constitutive transport vesicles from the TGN to the cell membrane (Kohler and Zahraoui, 2005).

3. Epithelial Morphogenesis into Tubes Tubular systems are generated using not only mechanisms for cell trafficking and polarization, but also mechanisms based on common elements of cell behavior associated with the three-dimensional (3D) architecture of the epithelial organs. Some of these mechanisms include cell migration to target sites, cell-fate diversification, and localization of specialized cells to different regions of the tube system. Tubular organs in vertebrates present a very complex organization with different cell types and localization (i.e., the kidney, the mammary, salivary and lachrymal glands). However, in simpler organisms, such as C. elegans and Drosophila, such epithelial organs consist of only few cell types, and a straightforward development program. In the past years many different cell models have been used to analyze the molecular and cellular events required to organize individual cells into 3D epithelial organs.

3.1. Model systems for studying tubulogenesis 3.1.1. In vitro models Several in vitro systems consisting on cultured epithelial cell lines grown in a layer of (or embedded in) extracellular matrix (ECM) have been developed to study the molecular and cellular events required to organize individual cells into epithelial organs. MDCK epithelial cell system is perhaps the best and most widely used in vitro model to investigate cell polarity during epithelial morphogenesis (Lubarsky and Krasnow, 2003; Martin-Belmonte and Mostov, 2008; Zegers et al., 2003). MDCK cells, which have properties of the kidney distal tubule and collecting duct, have been used for decades as a 2D model to study epithelial polarity and protein trafficking. However, since the filter support provides an overriding extrinsic cue to orient cell polarity, they represent a less appropriate model to analyze morphogenesis. By contrast, MDCK cells embedded in ECM form cysts, spherical monolayers enclosing a central fluid-filled lumen (Montesano et al., 1991), which have proven to be a very informative model system. Bissell, Brugge, and Werb labs have used 3D cultures of mammary cells for many years (Bissell et al., 2003; Mailleux et al., 2008; Sternlicht et al., 2006). Although, most mammary cell lines are not fully polarized, these 3D culture models illustrate how the microenvironment plays a critical role in regulating mammary

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tissue function and signaling. Experimental methods are provided to generate and manipulate 3D organotypic cultures to study the effect of matrix stiffness and matrix dimensionality on epithelial tissue morphology and signaling ( Johnson et al., 2007). Other in vitro systems use endothelial cells, a specialized form of epithelial cells, as a model (Bauer et al., 1992; Levin et al., 2007). The cell line EA hy926, derived from HUVECs cells, has proved to be a useful model for in vitro study of angiogenic processes. These cells plated on an ECM undergo a process of morphological reorganization leading to the formation of a complex network of cord, or tube-like structures. These events seem to resemble, in some aspects, an in vitro process of angiogenesis (Bauer et al., 1992). 3.1.2. In vivo models The most studied in vivo model systems for tubulogenesis include Drosophila trachea and salivary gland, zebrafish gut and vasculature, and single-cell kidney of C. elegans. The Drosophila trachea and salivary gland are genetic model systems for branched and unbranched tubes, respectively. Both organs begin as polarized epithelial placodes, which through coordinated cell shape changes, cell rearrangement, and cell migration form elongated tubes (Kerman et al., 2006). The last discoveries regarding the details of cell fate specification and tube formation in the two organs reveal significant conservation in the cellular and molecular events of tubulogenesis. In particular, in the morphogenesis of the Drosophila trachea, the control of cell invagination, migration, competition, and rearrangement is accompanied by the sequential secretion and resorption of proteins into the apical luminal space, a vital step in the elaboration of the trachea’s complex tubular networks (Affolter and Caussinus, 2008). A genetic approach using the zebrafish model has led to identification of mutations and molecules that are responsible for specification of endothelial cells, and differentiation of arterial and venous cells, as well as patterning of the dorsal aorta and intersegmental vessels. These studies highlight the unique utilities and benefits of the zebrafish system for studying development of embryonic blood vessels. In addition, the intestinal progenitor cells represent an excellent genetic model to analyze the complex process of intestinal morphogenesis, which involves interactions among multiple signaling pathways. Studies on morphogenesis are critical for elucidating the molecular basis of congenital gut defects and provide novel insights into intestinal oncogenic processes (Rubin, 2007; Zhong, 2005). Finally, although the nematode C. elegans has no kidney per se, it has proved to be an excellent model for studying renal-related issues, including tubulogenesis of the excretory canal, membrane transport and ion channel function, and human genetic diseases including autosomal dominant polycystic kidney disease (ADPKD) (Barr, 2005).

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Additionally, some labs have recently started to use ex vivo animal models (using mouse salivary and mammary glands mainly) that are generating abundant information about the molecular mechanism underlying tube formation and branching morphogenesis in superior vertebrates (Sternlicht et al., 2006).

3.2. Formation of epithelial tubes follows diverse patterns Strategies for making tubes can be classified according to whether the epithelial cells already have apical–basal polarity, or whether such polarity is acquired de novo in response to extracellular cues (Hogan and Kolodziej, 2002; Lubarsky and Krasnow, 2003). 3.2.1. Formation of tubes from already polarized epithelia Wrapping and budding are the two main mechanisms that have been described. Both require apical constriction, mediated by acto-myosin cables at the apex of the cells, and therefore are two mechanistically related processes (Lubarsky and Krasnow, 2003). Another common peculiarity is that in both mechanisms, tubes arise from a polarized epithelial sheet. The process for wrapping requires that a group of cells of an epithelial sheet undergoes apical constriction in a coordinated way, causing the bending of the epithelial sheet until the edges meets and seal to form a tubular structure with parallel orientation to the plane of the epithelium. Two examples of wrapping are the formation of the neural tube in vertebrates, and gastrulation in Drosophila (Dessaud et al., 2008; Leptin, 2005). The process for budding requires, similarly to wrapping, the invagination of the epithelial cells by apical constriction in a direction orthogonal to the epithelium plane, forming a new branched tube (Lubarsky and Krasnow, 2003). This mechanism is used during branching morphogenesis in many epithelial organs, such as salivary gland and the tracheal system in Drosophila (Kerman et al., 2006), as well as the formation of the mammalian lung (Metzger et al., 2008). 3.2.2. Formation of epithelial tubes from unpolarized groups of cells In a number of mechanisms described for epithelial tubulogenesis, tubes arise from clusters of cells or individual cells that are nonpolarized (Lubarsky and Krasnow, 2003). An important variable in these mechanisms for lumen formation is the requirement of cell death, which eliminates cells inside the luminal cavity. Experimental data obtained from different in vitro and in vivo models has characterized two basic mechanisms, cavitation and hollowing, for lumen clearance. In cavitation, the lumen is generated by apoptosis of cells in the middle of the structure, whereas in hollowing, the lumen is formed by membrane separation and/or repulsion. The mechanism of lumen formation in the 3D MDCK model can shift between hollowing

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and cavitation, depending on the degree of cell polarization (MartinBelmonte et al., 2008). However, the suppression of apoptosis by the expression of antiapoptotic factors, delays, but does not eliminate lumen formation, indicating that lumens eventually clear up by a caspase-independent death process, such as autophagy (Debnath et al., 2002; Mailleux et al., 2007).

4. De Novo Formation of Tubes: Conserved Pathways and Molecular Mechanisms Despite the diversity in the way cells assemble into tubes, there are many conserved morphogenetic processes. In this review, we have focused in summarizing the current findings obtained from the in vitro systems on the sequential steps implicated in the de novo formation of epithelial tubes. These include, at least, the orientation of the axis of polarity, and the symmetry breaking process at the level of the plasma membrane by the formation of the apical domain and the central lumen.

4.1. Initial cell–cell and cell–ECM interactions drive the formation of AJs and the orientation of the axis of polarity In a simple tube, all of the epithelial cells are oriented so that their apical surfaces face the central lumen (O’Brien et al., 2002). To build a tissue, the polarity of each cell must be coordinated. The first step for the organization of the epithelial architecture involves the concerted integration of polarizing cues from different sources. First, cells must sense their environment, including their position in relation to the surrounding cells. This can be mediated by direct interaction of cells with the ECM through a variety of receptors, such as integrins; and by the homotypic interactions of the cells with each other through cadherins to form cell junctions. The polarizing cues direct the cells on how they must organize spatially within the tissue, and how to orient the axis of polarity. To perform these functions, cells need to sense and modulate the interaction with the ECM; for instance, the stiffness and other mechanical and chemical properties of the ECM (Kass et al., 2007), and reorganize their cytoskeleton. The cadherin’s family of adhesion molecules controls the physical interactions between cells, and is particularly important for the dynamic regulation of adhesive contacts that is associated with diverse morphogenetic processes. In epithelial cells, E-cadherin forms the AJs, and facilitates the formation of the entire epithelial junctional complex (Gumbiner, 2005). In addition, during embryonic development, cadherins control the separation of distinct tissue layers, the formation of tissue boundaries, the changes in the shapes of tissues through cell rearrangements, and the conversions

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between different cell states (such as epithelium vs. mesenchyma) (Gumbiner, 2005). Furthermore, they are also essential in the maintenance of stable tissue organization, since they prevent the dissociation and spread of tumor cells (Cano et al., 2000). Cadherins associate with the actin cytoskeleton through the catenin family of proteins, and are associated with the cadherin-mediated cell adhesion at the AJs (Nelson, 2008). Although the association with the actin cytoskeleton is not required for the formation of the adhesive bond itself, it is required to produce the necessary force to generate the changes in cell shape and/or cell movement, to establish cell polarity and to organize the tissue. The orientation of epithelial polarity depends ultimately on the interaction of the cell with the surrounding ECM. Studies with epithelial cells grown in 3D, such as the 3D-MDCK system, have provided important information regarding how epithelial cells identify and interpret polarizing cues coming from the ECM, and how they respond by inducing signaling pathways that control the orientation of polarity. Rac1 was the first protein identified in this model to cause an inversion of polarity after disruption (O’Brien et al., 2001). Rac 1 (together with RhoA and Cdc42) belongs to the Rho-subfamily of small GTPases that function as chemical switches for biological processes (Etienne-Manneville and Hall, 2002; Jaffe and Hall, 2005). They cycle between GTP and GDP-bound states with different kinetics in specific subcellular compartments determined by their regulators: Rho GTP exchange factors (GEF) and Rho GTP activating proteins (GAP); and effectors (Bos et al., 2007). Further experiments demonstrated that the interaction of MDCK cells with collagen I causes activation of Rac1, since blockade of b1 integrin prevented activation of Rac1, and led to an inversion of orientation of cyst polarity, supporting the idea that Rac1 and b1 integrin are needed for normal orientation of polarity (Yu et al., 2005). Besides, activation of Rac1 induces polarized secretion and the assembly of laminin, which in turn activates Rac1 generating a positive feedback loop that reinforces the orientation signaling pathway (O’Brien et al., 2001; Yu et al., 2005). Recent findings have also demonstrated a role for RhoA, ROCK I, myosin II, PI3K, and protein kinase B in this polarity pathway (Liu et al., 2007b; Yu et al., 2008). How the orientation of epithelial polarity is determined is a key issue in biology. However, to differentiate this process from the establishment of polarity has been an arduous task for many years (Martin-Belmonte and Mostov, 2008; O’Brien et al., 2002). One reason is that in conventional epithelial cell culture experiments, cells are usually on an artificial substrate, which provides a strong cue for the cell to orient its apical surface pointing away from the support. An advantage of the 3D MDCK system is that the cells are in an isotropic environment lacking strong external cues to orient polarity and thus, are more sensitive to perturbations that alter the

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orientation of cell polarity. Additionally, a similar inversion of epithelial polarity has been detected in vivo during tumor progression (Adams et al., 2004). Interestingly, both integrins and cadherins seem to use similar signaling pathways to integrate the polarizing cues in the acquisition and maintenance of epithelial polarity. Previous work in vitro and in vivo has demonstrated important roles for small GTPsases in the establishment of the cadherin- and integrin-mediated adhesion. These include the Rho subfamily, mainly through Rac1 (O’Brien et al., 2001; Van Aelst and Symons, 2002; Yu et al., 2005), Arf GTPases and Rap1, a ras subfamily member (Fujita et al., 2006; Price et al., 2004; Rangarajan et al., 2003). In addition, integrins and cadherins have the ability to induce, through the PI3K, the generation of PtdIns(3,4,5)p3 (Kovacs et al., 2002; Velling et al., 2008), a phosphoinositide that controls the formation and identity of the basolateral plasma membrane (Gassama-Diagne et al., 2006) (discussed in detail below). Since GTPases and phosphoinositides regulate the cytoskeleton in different cell types and processes (Fukata et al., 2003), they could consequently mediate the connection of cadherins and integrins with the cellular cytoskeleton and other important processes associated with the formation of a polarized epithelia.

4.2. Vesicle trafficking, membrane separation, and lumen coalescence in lumen formation Once the epithelial cells have established apical–basolateral polarity and formed junctional complexes, the next step is to form the central lumen. The mechanisms by which this lumen is formed a key question in morphogenesis (Lubarsky and Krasnow, 2003). Emerging data from different models have concluded that, probably in all systems, epithelial cells create lumens following a series of events associated with membrane trafficking (Fig. 3.4) that include apical membrane biogenesis, transport of apical vesicles to the plasma membrane, secretion, and regulated expansion (Lubarsky and Krasnow, 2003; O’Brien et al., 2002). Lumen formation factors and other apical targeted proteins might be delivered to the nascent luminal surface by exocytosis of a specialized organelle, the vacuolar apical compartment (VAC), which is made of membranes that resemble the luminal plasma membrane. VACs are thought to be formed by endocytosis of a small portion of their external plasma membrane to create an internal vesicle containing some of the extracellular fluid (Fig. 3.4). These vacuoles fuse with each other and with the plasma membrane to produce small lumens between cells that expand to acquire the final size (Fig. 3.4). The existence of these VACs was characterized recently in vivo in the developing blood vessels of fish embryos (Kamei et al., 2006), although it could follow different morphogenesis programs (Blum et al., 2008). In epithelial cells, VACs seem to appear only in nonphysiological situations, either when the calcium levels are drastically

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Figure 3.4 Mechanism for de novo lumen formation in epithelia. Top, confocal sections of control and Cdc42-siRNA knock down MDCK cysts. Images of fixed cells in 3D were taken at the indicated times. Cysts are stained for an apical marker (gp135, red); adherent junction marker (b-catenin, green), and nuclei (blue). Arrowheads indicate accumulation of intracellular VACs close to the plasma membrane in MDCK cysts knock down for Cdc42. Scale bar 5 mm. Bottom, general mechanism for generation of lumens, including: (1) apical membrane biogenesis mediated by the endocytosis of apical proteins into VACs and vectorial transport of VACs to the plasma membrane, (2) fusion of VACs with the plasma membrane mediated by the Cdc42 dependent pathway, and (3) secretion, and regulated expansion.

reduced with chelators, such as EGTA (Vega-Salas et al., 1987), or when apical-vesicle delivery is delayed. In fact, the accumulation of apparent VACs was observed when Cdc42 was depleted in the 3D MDCK model, indicating a role for Cdc42 in the exocytosis of VACs to form the lumen (Martin-Belmonte et al., 2007). Confirming these observations, the expression of a dominant negative form of Cdc42, Cdc42N17, blocked capillary lumen formation in vitro (Bayless and Davis, 2002). Recent data has demonstrated a function for Cdc42 in the orientation of the mitotic spindle to position the apical surface during epithelial morphogenesis ( Jaffe et al., 2008), which suggest that Cdc42 must control different pathways during epithelial morphogenesis. In addition, other elements of the apical sorting machinery such as MAL/VIP17, Annexin 2, Annexin 13, Galectin-3, Syntaxin 3 and FAPP2, have been found to regulate lumen formation in the MDCK-3D model system (Martin-Belmonte et al., 2007; Torkko et al.,

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2008; Vieira et al., 2005). Rab11 and Rab8, two small GTPases implicated in membrane traffic, regulate exocytosis of apical proteins in vivo to form the lumen (Li et al., 2007; Sato et al., 2007). Epithelial secretion and endocytosis have also been required to drive airway maturation in the traqueal system (Behr et al., 2007; Tsarouhas et al., 2007). Finally, different proteins associated with vesicle transport in Drosophila photoreceptor cells have been characterized to have a role in lumen formation in morphogenesis (Beronja et al., 2005; Sang and Ready, 2002). A critical event for membrane separation and lumen formation is the presence of antiadhesive and/or repulsive factors in the apical surface to prevent the membrane from sticking. This may be mediated by secretion of large transmembrane glycoproteins or polysaccharides, which may induce membrane detachment by steric hindrance of cell–cell adhesion, and/or serve as a scaffold for the formation of the luminal space. For example, several groups showed that correct dilation of the tracheal tubes in Drosophila requires the formation of a transient luminal chitin-based matrix, which coordinates tube growth (Swanson and Beitel, 2006). Somewhat similarly, the Drosophila retina forms an epithelial lumen, the interrhabdomeral space, by the secretion of eyes shut, a protein closely related to the agrin and perlecan proteoglycan, into apical matrix of the luminal space. Eyes shut mutants fail to open the luminal cavity (Husain et al., 2006). In vertebrate development, renal glomerular epithelial cells (podocytes) undergo extensive morphologic changes necessary for creation of the glomerular filtration apparatus that include opening of intercellular urinary spaces. Podocalyxin, also known as gp135, a sialomucin, keeps the urinary space open by virtue of the physicochemical properties of its highly negatively charged ectodomain (Orlando et al., 2001; Takeda et al., 2000). This function for gp135/podocalyxin was confirmed recently in the 3D MDCK model (Meder et al., 2005). As mentioned before, membrane repulsion is a lumen formation event related to antiadhesive factors. Recent results have shown that Slit and its transmembrane receptor Robo play central roles in cardiac lumen morphogenesis functioning in autocrine signaling (Medioni et al., 2008; SantiagoMartinez et al., 2008). Slit–Robo signaling has an extensively studied function in repulsive neuronal axon guidance (Dickson and Gilestro, 2006). Why does lumen formation fail when Slit–Robo signaling is compromised? Medioni, Sanrtiago-Martinez, and co-workers have described that regulation of cell adhesion is a key factor, with Robo and E-cadherin having apparently opposing roles in lumen formation. Heart lumen formation is distinct from typical epithelial tubulogenesis mechanisms because the heart lumen is bounded by membranes that have basal rather than apical properties. In fact, lumens form as Slit–Robo signaling antagonizes E-cadherin-based adhesion specifically at the luminal domains of apposing cardioblast cells.

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4.3. The role of PtdIns in the definition of membrane identity Phosphoinositides have been implicated in nearly all aspects of cell physiology. Particularly, they function as general markers of membrane identity (Di Paolo and De Camilli, 2006). Diverse phosphoinositides are enriched in specific subcellular compartments, concentrated at the cytosolic surface of cellular membranes. The plasma membrane is enriched in phosphatidylinositol-4,5bisphosphate (PtdIns(4,5)p2), whereas PtdIns(3)p is found in endosomes and might specify the identity of various membranous compartments. Phosphatidylinositol-3,4,5-trisphosphate (PtdIns(3,4,5)p3) appears to be also associated with the plasma membrane and play conserved roles in the determination of cell polarity in very diverse cell types (Comer and Parent, 2007; Leslie et al., 2008). Furthermore, it has been proposed that the signaling mediated by (the balance of) PtdIns(4,5)p2 and PtdIns(3,4,5)p3 at the plasma membrane, together with the enzymes that regulate them, PTEN and PI3K, controls many of the most important cellular processes such as growth, polarity, motility, and proliferation (Carracedo and Pandolfi, 2008). PtdIns achieve direct signaling effects through the binding of their head groups to cytosolic domains of membrane proteins or cytosolic proteins. The PtdIns recruit the cytosolic proteins through phosphoinositide-binding modules, such as the pleckstrin homology domain (PH). Thus, they can regulate the function of integral membrane proteins, or recruit to the membrane cytoskeletal and signaling components (Di Paolo and De Camilli, 2006). 4.3.1. PtdIns 4,5 P2 is enriched at the apical domain and regulates the formation of the apical membrane and the lumen In mammalian epithelial cells, PtdIns(4,5)p2 is a key determinant of the apical surface, whereas PtdIns(3,4,5)p3 is a determinant of the basolateral surface (Gassama-Diagne et al., 2006; Martin-Belmonte and Mostov, 2007). In the 3D MDCK system, during the early stages of polarization, PtdIns(4,5)p2 becomes enriched at the apical membrane delimiting the newly formed lumen (Fig. 3.5). In contrast, PtdIns(3,4,5)p3 is exclusively localized to the basolateral membrane and excluded from the apical membrane (Fig. 3.5). The lipid phosphatase, PTEN (phosphatase and tensin homolog deleted on chromosome 10), which converts PtdIns(3,4,5)p3 to PtdIns(4,5)p2 becomes localized early to the apical domain, and its activity is required both for segregation of the two lipids, and for normal morphogenesis in different epithelia (Fig. 3.5) (Leslie et al., 2008). PtdIns(4,5)P2 functions for specific tethering of membrane binding proteins including cytoskeletal related proteins, such as vinculin, talin or ERMs, RhoGTPase activators, such as many Dbl family GEFs, and tethering complexes, such as annexin A2 complex and the exocyst (Liu et al., 2007a) and this may underlie the role of phosphoinositides in membrane identity. However, our understanding of how phosphoinositides are connected to cell polarity is still not complete.

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Figure 3.5 Cdc42 and phosphoinositides regulates the acquisition of apical– basolateral polarity during epithelial morphogenesis. PTEN activity mediates the enrichment of PtdIns(4,5)P2 (green) at the apical domain and restricts PtdIns(3,4,5)P3 (red) to the basolateral surface. The localization of PTEN to tight junctions is mediated by Par3, whereas its localization to the apical cortex could by mediated by NHERF. PI3K might localize to the adherent junctions (blue) and contribute to the stable presence of PtdIns(3,4,5)P3 at the basolateral membrane, which is necessary for the organization of this domain. PtdIns(4,5)P2, Annexin A2, and a specific GEF at the apical domain targets and activates Cdc42, which in turn activates Par6/aPKC and other effectors. Activated Cdc42 regulates the actin cytoskeleton (purple), which mediates the fusion of VACs with the plasma membrane to form the apical domain.

A possible explanation may involve the small GTPase Cdc42 and Par3. Cdc42 is a center molecule in different polarity processes in unicellular and multicellular organism (Etienne-Manneville, 2004). It is well known that PtdIns(3,4,5)P3 and PtdIns(4,5)P2 are required for the recruitment and activation of the Rho GTPases family to the plasma membrane, to modulate actin dynamics in various cell types (Etienne-Manneville and Hall, 2002; Yin and Janmey, 2003). For instance, targeting of PtdIns(4,5)P2 to the cell membrane leads to the recruitment and activation of Cdc42 in Xenopus eggs extracts. In cytoplasmic extracts from Xenopus eggs, PtdIns(4,5)P2 initiates an actin nucleation pathway that, synergistically with Cdc42, activates N-WASP, which in turn stimulates the actin-nucleating activity of the Arp2/3 complex (Rohatgi et al., 1999, 2000). In the 3D MDCK model, PtdIns(4,5)P2 induces the recruitment and activation of Cdc42 (MartinBelmonte et al., 2007). Cdc42 also binds to the Crib domain of Par6 and is necessary for correct localization of Par6/aPKC (Fig. 3.5), as well as for normal apico–basal polarization of Drosophila neuroblasts and epithelial cells (both in 3D MDCK and in Drosophila) (Atwood et al., 2007; Hutterer et al., 2004; Martin-Belmonte et al., 2007). Furthermore, recent

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results in Drosophila neuroblasts have shown that Cdc42 localizes Par6/ aPKC at the apical cortex in a Par3-dependent manner, indicating that Par3 is upstream of Cdc42 in these cells (Atwood et al., 2007), and upstream of Par6/aPKC in embryonic epithelial cells (Harris and Peifer, 2005). Par3 also plays an important role in connecting phosphoinositides and cell polarity. Par3 contains multiple PDZ domains and binds via its second PDZ domain to most species of phosphoinositides, including both PtdIns (4,5)P2 and PtdIns(3,4,5)P3 (Wu et al., 2007). This binding is needed for the recruitment of Par3 to the plasma membrane and for polarization of MDCK cells, at least in 2D culture. However, as Par3 binds to both of these lipids with nearly equal affinity, this lipid binding alone would not appear to explain the localization of Par3. In Drosophila epithelial cells, Bazooka/Par3 binds to PTEN and localizes it to the cell–cell junctions (von Stein et al., 2005). Although this interaction has also been described in vitro in MDCK cells (Fig. 3.5) (Wu et al., 2007), PTEN is also associated with the apical membrane in mammalian cells forming 3D structures (Martin-Belmonte et al., 2007). Therefore, more work will be needed to address what localizes PTEN in mammalian cells. Interestingly, PTEN interacts directly with the NHERF1 adaptor protein (Naþ/Hþ exchanger regulatory factor required to organize complexes at the apical membranes of polarized epithelial cells) through the PDZ motif of PTEN and the PDZ1 domain of NHERF1 (Morales et al., 2007). Additionally, Par3 malfunction has proved to induce defects in central lumen formation in vitro in the 3D-MDCK model system (Hurd et al., 2003a), and also in vivo in cardiac cyst development in mice (Hirose et al., 2006). 4.3.2. PtdIns 3,4,5 P3 is restricted to, and defines, the basolateral domain PtdIns(3,4,5)p3 is emerging as a spatial landmark of specialized regions of the plasma membrane, such as the axon growth cone, phagocytic cup, leading edge of chemotaxing cells and can even determine the orientation of the mitotic spindle (Toyoshima et al., 2007). Interestingly, addition of exogenous PtdIns(3,4,5)p3 to the apical surface of filter-grown MDCK cells causes the transformation of the apical surface into basolateral within a few minutes. These experiments show that phosphoinositides are indeed molecular determinants of the identity of the apical and basolateral surfaces even in 2D epithelial cultures. In further support of the hypothesis that PtdIns(3,4,5)p3 specifies the basolateral surface, growth of MDCK cells in the presence of low concentrations of inhibitors of phosphatidylinositol-3 kinase (PI3K) reduced the size of the lateral membrane and height of the cells. Remarkably, binding of Pseudomonas aeruginosa, a bacterial pathogen that infects epithelia through the basolateral surface, to the apical surface leads to localized accumulation of PI3K, PtdIns(3,4,5)p3, and basolateral proteins at the apical surface, similar to the experimental addition of PtdIns

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(3,4,5)p3 to the apical surface. In essence, the bacteria transform the apical surface into basolateral, thereby enabling invasion of the host cell (Kierbel et al., 2005, 2007). However, the molecular details that define these relationships in different cells and tissues are probably distinct. In Drosophila photoreceptor epithelial cells, the recruitment of PTEN to cell–cell junctions, spatially restricts PtdIns(3,4,5)p3 to a the rhabdomere, a highly specialized region of the apical domain of these cells (Pinal et al., 2006). This result highlights that PtdIns(3,4,5)p3 can have a flexible role in determining specialized regions of a wide variety of cell types. There are other situations where PtdIns(3,4,5)p3 is involved in remodeling a specialized domain of the cell surface. For instance, stimulation of kidney cells by insulin binding to the basolateral surface leads to accumulation of PtdIns(3,4,5)p3 at the apical surface, followed by insertion of an epithelial sodium channel into the apical membrane (Blazer-Yost et al., 2004). Together, PtdIns(4,5)p2 and PtdIns (3,4,5)p3 seem to play essential roles in epithelial polarity and plasma membrane identity.

5. Epithelial Polarity and Disease There is growing evidence that a wide array of mutations in epithelial polarity genes is implicated in human disease (both monogenic and polygenic). Some of these genes have house-keeping functions in many cell types and their disruption in knockout mice models results in embryonic lethality. Many are found mutated with altered structure or expression, and are cause of diverse pathologies. Interestingly, many epithelial genes are related to a higher risk of developing cancer, and some of them are well known proto-oncogenes and tumor suppressors. In this review, we first address the relationship between epithelial polarity and cancer, and then we focus on mutations that are involved in syndromes caused by trafficking alterations. Finally, we address cytoskeletal, phosphoinositide and GTPase related disorders.

5.1. Cell polarity and cancer A vast majority of malignant human cancers are originated in epithelial tissues. Epithelial neoplastic tissues are characterized by loss of cell–cell junctions, loss of epithelial apico-basal polarity, and an increase in proliferation rates. Cells evade apoptosis and cancer progresses, as they acquire the mesenchymal phenotype, invading the inner tissues, a process called EMT (Hanahan and Weinberg, 2000). The regulation of cell proliferation and its related genes have been extensively studied. However, the relationship between polarity and proliferation is just recently starting to be unveiled. Indeed, current evidence shows that many mutated genes identified in

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aggressive cancers, are related to cell polarity establishment and maintenance, suggesting that cell polarity could act as a noncanonical tumor suppressor (Lee and Vasioukhin, 2008). Cancer cells that have lost their polarity regulation are more prone to cause metastasis, which is the spreading of cancer cells throughout multiple body tissues, and a hallmark of advanced cancer (Dow and Humbert, 2007). The key factor in cell–cell junction formation in epithelia is E-cadherin (Takeichi, 1991). Most epithelial cancer cells either harbor mutations in E-cadherin genes or lose E-cadherin expression along the way towards EMT (Thiery and Sleeman, 2006). Induced loss of E-cadherin in intestinal tissues in mice either increase tumor migration or proliferation and mutations in CDH1 are normally found in stomach, pancreatic, and other epithelial cancers (Conacci-Sorrell et al., 2002; Perl et al., 1998). The Scribble and PAR complexes are both implicated in human cancer (Fig. 3.6). The members of the Scribble complex (Lgl, Scribble, and Dlg) are neoplastic tumor suppressors in Drosophila larvae, where their mutation causes overgrowth of imaginal disc cells and loss of cell polarity and tissue architecture, larvae dying before reaching pupation (Bilder, 2004; Bilder et al., 2000; Gateff, 1978; Peifer, 2000). Moreover, Lgl and Dlg loss causes metastasis when imaginal disc cells are injected in adult hosts (Woodhouse et al., 1998). In mammals, there are two homologous Lgl genes, Lgl1 and Lgl2. Lgl1 knockout mice show severe brain dysplasia and loss of polarity in neuroepithelial cells, but the expression of a homolog gene, Lgl2, probably provides a functional backup in the rest of tissues (Klezovitch et al., 2004). Mammalian Lgl, Dlg, and Scrib expression loss is commonly found in a wide array of epithelial cancers (lung, prostate, skin, breast and colon cancers, and Human Papillomavirus (HPV)-induced cervical cancers) (Humbert et al., 2003). Studies carried out using cells from tumour samples have shown that overexpression of Lgl and Dlg are capable of phenotype attenuation and restoring cell polarity (Kuphal et al., 2006; Massimi et al., 2004). In aggressive HPV strains, HPV 16 and 18, E6 oncoprotein is capable of targeting the Scrib complex for proteolytic degradation, a condition that correlates with invasiveness increase and progression of the cervical tumor (Massimi et al., 2004; Thomas et al., 2005). Finally, Lgl, Dlg, and Scrib mutations increase the metastatic behavior of oncogenic Ras-mediated cancers and Ras oncogenic activation increases E6-mediated transformation, synergizing along with Scrib and Dlg proteolytic downregulation (Fig. 3.6) (Brumby and Richardson, 2003; Storey and Banks, 1993). The PAR–aPKC complex is the main regulator of apico-basal polarity at the apical pole of the cell, where it phosphorylates and inactivates Lgl (Betschinger et al., 2003; Plant et al., 2003). In Drosophila, activation of aPKC promotes tumorigenesis in lgl mutants and constitutively active forms

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PIP3

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Figure 3.6 Epithelial cell polarity and cancer. Mechanisms and complexes implicated in epithelial cell polarity that promote tumorigenesis when become deregulated. PTEN mutations cause PIP3 accumulation, leading to upraised survival signaling via Akt activation. Annexin 2 (Anx2) expression is abnormal in breast, brain, pancreatic, and prostatic tumors. Downstream of Anx2 and Cdc42, PAR complex signaling defects are related to cancer through distinct manners. Atypical PKC (aPKC) is overexpressed in many human cancers, and constitutively active forms cause overgrowth in D. melanogaster imaginal discs. The Scrib complex proteins behave like tumor suppressors and are targeted for degradation by E6 oncoprotein, promoting Ras-mediated oncogenesis. Par3 is the target of oncogenic ErbB2, which disrupts tight junction signaling by inhibiting Par3 binding to the PAR complex. Tumor suppressor VHL protein regulates PAR complex activation through targeting Par6 for proteolytic degradation. Finally, oncogenic TGFb receptor mediates Par6 binding to Smurf1 E3 ubiquitin-ligase, inducing Rho GTPase degradation and depolimerization of the actin belt that secures polarized epithelial architecture, promoting EMT.

of aPKC cause epithelial polarity loss (Fig. 3.6) (Lee et al., 2006). Lgl mutant phenotypes can be rescued inactivating aPKC, suggesting aPKC deregulation could have a causal role in those phenotypes (Rolls et al., 2003). The main isoform of atypical PKC in mammals, aPKC-i has also been found amplified in human cancers (ovarian and nonsmall-cell lung cancers) and aPKC-i activity is needed for xenograft tumour development and anchorage-independent cell growth (Eder et al., 2005; Regala et al., 2005; Zhang et al., 2006a). Less studied aPKC-z is also hyperactivated in squamous-cell carcinoma of the head and neck, where it mediates EGF-induced MAPK activation, and in glioblastoma cell lines (Cohen

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et al., 2006; Donson et al., 2000). Par6 is a target for the serine kinase activity of TGFb-Receptor, which promotes EMT during both development and tumor progression (Fig. 3.6). Par6 phosphorylation results in Smurf1 E3ubiquitin ligase recruitment and specific degradation of RhoA at the apical pole, destabilizing the actin belt that provides the scaffold for apical junction complexes (Ozdamar et al., 2005). Another signaling receptor, ErbB2, overexpressed or amplified in the 25–30% of breast cancers, disrupts cell polarity and provides antiapoptotic tumour protection by displacing Par3 from the PAR complex (Fig. 3.6) (Aranda et al., 2006; Hynes and Lane, 2005). The von Hippel–Lindau protein (VHL) is an E3-ubiquitin ligase that is mutated in the hereditary clear cell renal cancer syndrome that carries the same name. VHL functions as a tumour suppressor, controlling aPKC activity through signaling Par6 for degradation, and its expression loss correlates with an increase in aPKC levels, affecting cell polarity and favoring cancer progression (Fig. 3.6). PTEN (Phosphatase and Tensin homolog deleted on chromosome 10) is another protein mutated in multiple epithelial cancers, implicated both in cell polarity and cell proliferation, and it is the disease gene in the hereditary Cowden cancer syndrome (Di Cristofano and Pandolfi, 2000). The main dysfunction associated to PTEN mutations is PI3K pathway deregulation, leading to sufficient and signal-independent cell growth and survival (Fig. 3.6) (Lemmon, 2008). As noted before, recent studies have related PTEN function with cell polarity development in MDCK cysts. The dual function of a tumor suppressor, acting both in cell polarity development and proliferation inhibition, is an important example of how both processes are intricately related. We can only speculate whether or not PTEN mutations promote cancer through its polarity-related functions. More studies using downstream effectors and the elucidation of a complete pathway for Cdc42 and Par6–aPKC apical activation remain to be key issues for answering these questions. PAR complex regulators and effectors are also partly responsible for effects in cancer disease. The main GTPase activator for the PAR complex, Cdc42, is frequently downregulated upon Ras hyperactivation, and loss of Cdc42 cooperates with oncogenic Ras in cells expressing Ras activated mutants, allowing tumorigenesis in Drosophila imaginal discs (Fig. 3.6) (Sahai et al., 2001). PAR complex effector PAR4/LKB1 is mutated in the heritable PJS (Hemminki et al., 1998). LKB1 mutations in PJS do not impair the kinase activity, but instead reduce the ability to promote cell polarization in gastrointestinal cells (Alessi et al., 2006). The recent discovery of AMPK as a cell polarity regulator (by means of actomyosin activation through rMLC phosphorylation) is another important example of the coordinated regulation that controls cell polarity and proliferation (Forcet and Billaud, 2007; Lee et al., 2007; Williams and Brenman, 2008).

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Annexins present an important role in human cancer progression. Most genome-wide cancer expression assays have pointed out at least some annexins as genes differentially regulated in neoplasia (Hayes and Moss, 2004). Prostate cancer cells downregulate Annexin A2 during cancer progression and Annexin A1 seem to have reduced level of expression in squamous carcinomas (Fig. 3.6) (Liu et al., 2003; Paweletz et al., 2000; Vishwanatha et al., 2004). In addition, Annexin A2 is overexpressed in pancreatic, breast and brain cancer, and its upregulation could be the cause of increased cancer angiogenesis to promote metastasis (Huang et al., 2008; Liu et al., 2003; Nygaard et al., 1998; Sharma et al., 2006; Vishwanatha et al., 1993). There is a constantly growing body of evidence supporting the cancer stem cell theory (Cho and Clarke, 2008; Clarke and Fuller, 2006). Stem cells are present in many adult tissues and represent a source for the renewal of epithelial cells in the skin, the gut, and the blood. This theory proposes that certain cells in the tumor behave like stem cells, and they could also be the key in the origin of cancer. Adult stem cells normally undergo asymmetric cell division, a process that allows them to provide differentiated progenitor cells while retaining their own proliferative capacity (Knoblich, 2008). The mechanism requires polarization of fate-determinants and is driven by cell polarity regulators, like the PAR and Scrib complex (Wodarz and Nathke, 2007). When asymmetric division goes wrong, stem cells divide into equivalent cells, both of which retain their renewal potential. Recent advances propose that this process could be at the heart of tumorigenesis (Morrison and Kimble, 2006; Wodarz and Gonzalez, 2006). Cancer stem cells (CSCs) have been isolated from many tumors in mammals (Dick, 2003; Lee et al., 2008; O’Brien et al., 2007; Singh et al., 2004; Wang and Dick, 2005). These immortal, undifferentiated, progenitors are somewhat similar to mutant Drosophila neuroblasts, which are unable to divide asymmetrically. Although mutations in most tumorigenic polarity genes in Drosophila do not cause any phenotype in most tissues in mammals, probably because of functionally redundant paralogues, there are well known cancer-related functions for many genes that regulate spindle orientation and the mitotic checkpoint. Aurora A and Polo are two kinases that are implicated in the spindle assembly checkpoint, and, when mutated or deleted, cells fail to segregate chromosomes correctly (Malumbres and Barbacid, 2007). As a result, daughter cells inherit unstable and abnormal genomes, where some genes may become deleted or amplified, predisposing for carcinogenesis. In higher eukaryotes, cells are embedded into three dimensional environments, with multiple mechanic and chemical interactions with their milieu, organizing with specific tissue architecture (Bissell et al., 2003). The maintenance of this microenvironment has proven to be essential to counteract the proliferative and invasive properties of cancers, as well as to

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keep cells differentiated (Yamada and Cukierman, 2007). Original studies in oncogenesis resulted from comparison between in vivo and in vitro tumorigenic cell lines. For example, Ras-mediated or oncovirus-mediated transformation is evident in cultured cells, but is insufficient to generate tumors by itself in a complete organism (Dolberg and Bissell, 1984; Frame and Balmain, 2000). Importantly, wild-type cells behaved in a completely different manner when they grew together with mutated cells in a tissue or in a culture plate. In culture, cancer cells seem to influence a tumor-like behavior on wild-type cells, while this is rarely seen in a complete organism where tumors do not normally change the phenotype of a whole tissue, but rather evolve clonally. In addition, some epithelial cancers behave differently in 2D versus 3D cultures. Single oncogenes that normally had effect in two dimensional cultures were unable to produce the same effect in 3D cultures (Partanen et al., 2007). A fundamental signal is provided to the cells through ECM interaction with integrins (Gumbiner, 1996). In mammary gland cells cultured in 3D, disruption of b1-integrin signaling (using blocking antibodies) was sufficient to attenuate ErbB2-mediated transformation and invasion (Weaver and Roskelley, 1997; Weaver et al., 1997). It was proposed that in mammary gland cancer integrin signaling could promote cell proliferation and invasion, through the Rac1 pathway and that it could protect epithelial cells from apoptotic cell death. Indeed, anoikis is a type of cell death associated with lack of ECM signaling (Zahir and Weaver, 2004). However, as stated earlier, integrin signaling in the 3D-MDCK model was proven essential for promoting adequate orientation of cell polarity (O’Brien et al., 2001). Recently, it was also proven that polarity could regulate apoptotic clearance of epithelial lumen, and that disruption of Cdc42 activated polarity signaling results in increase of apoptosis in developing cysts (Martin-Belmonte et al., 2008). We propose that apoptotic cell death could be activated by loss of polarity, as a tumor-suppressor mechanism that would prevent abnormal growth of deregulated cells, although the underlying mechanisms remain unclear.

5.2. Trafficking disorders As discussed previously, protein trafficking routes are among the most complex mechanisms in cellular physiology. Complex machinery including GTPases (Rab proteins), adaptor complexes (APs), and tethering and docking complexes (Annexins) have proven to be related to some extent with human disease (Fig. 3.7) (Aridor and Hannan, 2000, 2002). 5.2.1. Defects in protein sorting signals A wide number of monogenic diseases are caused by point gene mutations that distort the sorting signal, resulting in the incorrect distribution of a specific protein and originating the disease (Stein et al., 2002).

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Rab8

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Figure 3.7 Trafficking and cytoskeletal disorders. Defects in the correct sorting and trafficking of vesicles cause a wide array of human pathologies (in red). Alterations in peptidic sorting signals are responsible for CFTR mislocalization and defects in ion transport (Cystic Fibrosis, CF). Defective trafficking machinery (such as Myo5B or Rab8) originates dysfunctional apical membrane domains and apically targeted vesicle accumulation (Microvillus Inclusion Disease, MVID). Abnormally high levels of Rab activation are common in cancer disease, and defects in Rab targeting to vesicle membranes (caused by REP1 mutations) originate specific problems in specialized epithelia (such as retinal choroideremia). Vesicle formation is mediated by adaptor protein complexes, such as AP-3 which causes defects in lysosomal maturation in the Hermansky–Pudlack Syndrome (HPS). Finally, Rho family GTPases activation is mediated by Rho GEFs, which are overexpressed or gain-of-function mutated in many cancers (i.e., LARG, TIAM, and Dbl GEFs). Mutations in Rho GTPase effectors, such as Diaphanous 1 formin (Dia1) or WAS protein (WASP), inhibit actin polymerization and are cause for nonsyndromic deafness in humans and the Wiskott–Aldrich syndrome disease, respectively.

Cystic fibrosis (CF) is an autosomal recessive disease that affects chloride transport in epithelia and that arises through point mutations in a c-AMP sensitive chloride channel, CF transmembrane conductance regulator (CFTR) (Riordan, 2008). The protein is located to the apical membrane compartment in renal cells. Its cytoplasmic tail contains a PDZ binding motif that interacts with CFTR-associated ligand (CAL) (Moyer et al., 1999), and C-terminal apical trafficking motifs that interact with Naþ/Hþ exchange

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regulatory factor (NHERF) (Milewski et al., 2001). Mutations in the cytoplasmic tail result in loss of CFTR apical localization which in turn originates defective chloride transport and CF (Fig. 3.7) (Ameen et al., 2007). Recently, it was shown that myosin VI knock-out mice have a gastrointestinal CFTR phenotype, with loss of enterocytic brush border, implicating myosin VI in CFTR trafficking in vivo (Ameen and Apodaca, 2007). Wilson disease is a monogenic autosomal recessive disease of copper metabolism that affects hepatic copper excretion. The gene, ATP7B, codifies a cell surface receptor that forms complexes with copper and is transported apically to release the cation at the lumen of the epithelium (Forbes and Cox, 2000). High levels of copper regulate ATP7B trafficking to the apical membrane. ATP7B mutations affect either cation binding or ATP7B trafficking to late endosomes (LE), causing the liver to accumulate copper and ultimately producing metal toxicosis (Morgan et al., 2004). Familial hypercholesterolemia is an autosomal genetic disease caused by a failure in LDL removal from plasma by hepatocytes. The most famous LDLR mutation is the autosomal recessive mutation described by Brown and Goldstein, which alters LDLR clathrin mediated endocytosis (Brown and Goldstein, 1976; Davis et al., 1986). However, autosomal dominant familial hypercholesterolemia mutations also map to the LDL receptor gene (LDLR) and are of relevance to this review. LDLR is normally expressed and localized to the sinusoidal membrane through two tyrosine-based basolateral sorting signals (Matter et al., 1992). Point mutations in these signals translate into a missorting of LDLR to the apical membrane compartment and the inability of hepatocytes to eliminate LDL from plasma, leading to LDL accumulation (Koivisto et al., 2001). The consequences rise as heart disease and atherosclerosis because of LDL deposition, plaque formation, and inflammation at the arteries. 5.2.2. Defects in sorting signal recognition and vesicle formation—Adaptor protein complexes Deletion or disruption of either AP1 or AP2 is absolutely lethal and mutations are rare among population. While mammalian mutations in AP-4 have not been found, the model for a human disease, called Hermansky–Pudlak syndrome (HPS), resulted from mice which had mutations AP-3 subunits (Fig. 3.7), either in AP3D1 (Delta-adaptin) or in AP3B1 (b3A) (Feng et al., 1999). The AP3B1 mutations were finally discovered in a small number of human HPS patients (Dell’Angelica et al., 1999). The autosomal recessive syndrome is related to deficiency in protein sorting to lysosomal and lysosomal-derived organelles (such as melanosomes and platelet granules) and human patients suffer from oculocutaneous albinism and platelet storage disease, with variable symptoms arising in subphenotypes (HPS1-8). Other murine HPS genes have also been related with cell polarity and trafficking,

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such as Vps33a (which sorts proteins to the vacuole in yeast) and RabGGTase (the enzyme catalyzing geranyl-geranylation of RabGTPases) (Wei, 2006). 5.2.3. Defects in Rab GTPases RabGTPases regulate vesicle transport and fusion spatially and temporally in a large number of cellular pathways, which are normally orchestrated by a specific member of the family. Many Rab and Rab-associated proteins have been implicated in human disease, ranging from neuropathies to cancer (Seabra et al., 2002). Choroideremia is an X-linked retinal degeneration disease which ultimately leads to inoperable blindness. It is caused by mutations in the Rab escorting protein gene REP1 (Fig. 3.7) (Andres et al., 1993; Rak et al., 2004; Seabra et al., 1992). Another escorting protein, REP2 seems to be able to compensate REP1 deficiency in the rest of tissues, thus affecting only the retinal epithelium where both of them seem to be needed (Cremers et al., 1994). TSC is an autosomal dominant disease characterized by formation of hamartomas, renal failure, and mental retardation. Mutations in two proteins were discovered in TSC suffering patients and were named hamartin (TSC1) and tuberin (TSC2) (Crino et al., 2006). Both genes have tumor suppressor activity and their products form a complex that acts downstream of the insulin receptor signaling pathway in which they are phosphorylated and inactivated by Akt, halting mTOR signaling (Gao et al., 2002; Inoki et al., 2002; Potter et al., 2002). The complex has Rab GAP activity, specific for Rab5 and Rap1, implicated in endocytosis, and Rheb GAP activity, thus shutting down the mTOR pathway (Fig. 3.7) (Wienecke et al., 1995; Zhang et al., 2003). The complex also regulates polycystin-1 trafficking, and in patients with tuberin mutations, polycystin-1 mislocalizes to the Golgi, with consequent cystic disease (Kleymenova et al., 2001). Polycystin-1 is the most vastly mutated gene in renal ciliopathies (discussed later) and the fact that tuberin mutations impede its normal trafficking accounts at least in part for the renal phenotype of the disease. Not only disruption of RabGTPase function but also abnormal RabGTPase activation is an important cause of human disease (Fig. 3.7). Rab5a and Rab7 are overactivated in Thyroid autonomus adenomas, and Rab7 is also overexpressed in atherogenesis (Croizet-Berger et al., 2002; Kim et al., 2002). Rab1a, Rab4, and Rab6 are upregulated in Beta2-AR mutant mice (which are human cardiomyopathy models) (Wu et al., 2001). Rab25 is also amplified in ovarian and breast cancer cells and a novel prostate cancer mutated gene, PRC17 has shown to be able to effect as an upregulated Rab GAP for Rap1 and Rab5 in human prostate cancer (Caswell et al., 2007; Cheng et al., 2004; Pei et al., 2002).

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Rab8, a RabGTPase implicated in basolateral protein sorting and adherens junction assembly, has been related to retinal degeneration and intestinal microvillus inclusion disease (MID) (discussed later).

5.3. Cytoskeletal and phosphoinositide related disorders 5.3.1. Defects in Rho family GTPases and Rho regulators The role of GTPases in cancer has been reviewed extensively (Vega and Ridley, 2008). Many GTPase pathways are linked directly to cell proliferation, polarity and migration, and their alterations are intimately linked to cancer progression and metastasis. Results from in vivo GTPase studies are beginning to arise (Heasman and Ridley, 2008). Gene targeting and disruption in mice models have proven both Rac1 and Cdc42 knock-out mice are lethal and die during development, at embryonic stages. However, mutations and diseases related to RhoGTPase defects are usually linked to their regulators, GAPs and GEFs, and effector genes, which are more specific in function (temporally and spatially) and whose mutations do not affect a wide number of processes (Fig. 3.7) (Chen et al., 2000; Sugihara et al., 1998; Wang and Zheng, 2007). Some RhoGEFs are well characterized oncoproteins in vitro (Cerione and Zheng, 1996). Many activating mutations have been found in Dbl family of GEFs during analysis of tumour samples including LARG (leukemia-associated Rho GEF), TIAM (T cell lymphoma invasiveness and metastasis), and Dbl (Diffuse B-cell lymphoma). Dbl was originally isolated as an oncogene (diffuse B-cell lymphoma), while LARG was isolated as a LARG-MLL gene fusion in acute myeloid leukemia, and TIAM was found in a screening for invasiveness related genes using T cells. The TIAM specific activation of Rac and its upregulation and mislocalization in aggressive cancerous cells is the hallmark for the discovery of GTPase function in cancer metastasis and tissue colonization. Faciogenital dysplasia 1 (FGD1) is a Dbl family RhoGEF, specific for Cdc42 activation (Olson et al., 1996; Pasteris et al., 1994; Zheng et al., 1996). It is the disease gene for the hereditary Aarskog–Scott syndrome, characterized by facial and urogenital malformations. FGD1 is recruited to membranes where actin functions must be carried out, by means of its PH domain. Once there, it activates Cdc42 to regulate the cortical actin cytoskeleton. Typical mutations implicate gene translocation or nonsense mutations. However, a subset of point mutations affects the PH domain (Orrico et al., 2000). The phosphoinositide specificity of its PH domain has not been assayed, but this fact signifies the importance of GEF membrane-binding domains in vivo for efficient GTPase function and a role for PH domains in disease. WASP is the product of the gene mutated in Wiskott–Aldrich syndrome (WAS), a hematopoietic disease that is characterized by recurring infections (Fig. 3.7) (Aldrich et al., 1954; Derry et al., 1994). Insights into its function

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were provided by studying its homologue N-WASP, originally isolated from brain tissue. WASP is a Cdc42 effector and stimulates Arp2/3 dependent actin polymerization, supporting the cortical matrix, and the cellular morphology (Symons et al., 1996). WASP missense loss-of-function mutations are the most frequent cause of WAS, originating cytoskeletal abnormalities in hematopoietic linage cells and immune system defects. Another GTPase effector related with actin polymerization is Diaphanous 1 (DFNA1, Dia1), homolog of D. melanogaster gene (Lynch et al., 1997). Its autosomal dominant mutation causes nonsyndromic deafness in humans, an inner ear dysplasia (Fig. 3.7). Mammalian Dia1 (mDia1) belongs to the Diaphanous-related formin (DRF) superfamily (Goode and Eck, 2007). It is activated specifically by Rho binding, upon which it linearly polymerizes actin and works in concert with ROCK inducing stress fibers, and aligning actin and microtubules through its microtubule binding domain. The mutations affect the actin cytoskeleton of hair cells in the cochleosaccular region of the inner ear, causing morphological abnormalities and deafness (Muller and Littlewood-Evans, 2001). 5.3.2. Microvillus inclusion disease Microvillus inclusion disease (MID), or Davidson’s disease, is a familial enteropathy with autosomal recessive inheritance (Cutz et al., 1989). Enterocyte and colonocyte differentiation is impeded and cells develop abnormal epithelial polarity (Ruemmele et al., 2006). Intestinal cells loose their brush border apical structure, and apical proteins and microvilli appear in intracellular vacuoles (Ameen and Salas, 2000). It is an extremely rare disease (with only about 30 cases diagnosed since its discovery in 1978). The interest for the disease became of notice when the VACs were first proposed as intermediates in epithelial polarity development and lumen morphogenesis (see Section 4.2) (Vega-Salas et al., 1988). VACs resembled intracellular microvilliar structures observed in MID and suddenly it became clear that MID patients could carry mutations in some of the genes involved in VAC formation and/or exocytosis (Fig. 3.7). Initial experiments were directed towards the actin and microtubule cytoskeleton and cytoskeleton binding proteins. MID patients present decreased expression of actin, vinculin, and myosin. Moreover, disruption of actin or tubulin cytoskeleton using drugs and inhibitors proved to mimic the MID phenotype. However, the search for the MID gene proved to be extremely difficult, due to the low incidence of the disease. Recent observations in the field have shown that Rab8 knockout mice display the MID phenotype (Sato et al., 2007). Mutations in Rab8 have not been proven to correlate directly with MID in humans, but at least one patient shows reduced expression of the gene. Also, genetic studies in a MID family have revealed a series of mutations in MYO5B, a gene encoding a myosin V motor protein isoform. The mutations cause alterations in transferrin receptor trafficking and disrupt epithelial cell polarity, along with

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microvilliar inclusions (Muller et al., 2008). Interestingly, Rab 8 and myosin V interact during endosomal transport (Roland et al., 2007). These studies have demonstrated a crucial role for myosin V and Rab8 in basolateral and apical trafficking and their relationship with MID (Fig. 3.7). Although these are promising results, not all MID patients carry mutations in MYO5B or Rab8, indicating that the phenotype might be a common consequence of diverse alterations in apical trafficking machinery. 5.3.3. Defects in phosphoinositide metabolism As emphasized earlier, phosphoinositides are implicated in many aspects of polarized cell physiology. The enzymatic network that determines the concentrations of each phospholipid subspecies is under tight regulation, by means of diverse signaling routes, and results in accumulation of specific lipids in a specific temporal and spatial frame in the cell. Mutations in PTEN are perhaps the most well known and studied, because of their implication in cell transformation and cancer (discussed above). However, a wider role for phosphoinositide signaling in trafficking and human disease is starting to emerge (Halstead et al., 2005; Nicot and Laporte, 2008). Other mutations in phosphatases include SHIP1, SHIP2, myotubularins, and OCRL1. OCRL1 is the phosphatase that converts PtdIns(4,5)P2 into PtdIns(4)P, a fundamental signaling lipid for lysosomal sorting from the TGN (Lowe, 2005). Mutations in this gene are found in patients suffering the Oculocerebrorenal syndrome of Lowe. The syndrome affects epithelial lens cells, renal tubule cells, and brain cells, causing bilateral congenital cataracts, Fanconi syndrome, and neurodegenerative retardation. In kidney tubule cells, OCRL1 mutations cause PtdIns(4,5)P2 to accumulate at the TGN (where OCRL1 is normally localized, and probably exerting its function) (Choudhury et al., 2005; Zhang et al., 1998). As a result, lysosomal proteins are missorted to the apical region, affecting the polarized epithelial phenotype and normal tissue integrity, and highlighting the importance of phosphoinositides in membrane trafficking. However, as we explained before, PtdIns(4,5)P2 domains are also a platform for linking lipidic membranes to cytoskeleton. OCRL1 cells lose their ability to form stress fibers and other F-actin based structures, and F-actin regulators gelsolin and alfa-actinin are abnormally distributed. Also, OCRL1 is recruited to membrane ruffles by means of Rac activation in response to growth factor signaling. Recently, it has been found that OCRL1 interacts with diverse RabGTPases, including Rab1, Rab5, and Rab6 (Hyvola et al., 2006). Rab5 effector APPL1 recruits OCRL1 to clathrin coated vesicles during endocytosis through an ASH-RhoGAP-like domain present in OCRL1 (Erdmann et al., 2007). Some OCRL1 mutations locate in this domain and impede normal recruitment, affecting normal membrane to TGN retrograde transport (McCrea et al., 2008). Studies using Ocrl1 knockout mice have proven a functional overlapping between this gene and its homologue Inpp5b,

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explaining the absence of phenotype ( Janne et al., 1998). In summary, actin polymerization, lysosomal sorting, and receptor endocytosis all result from PIP2 metabolism impairment.

5.4. Cystic diseases of the kidney Cystic diseases of the kidney (CDKs) are a group of monogenic diseases characterized by the appearance of fluid-filled cysts and end-stage renal failure (Hildebrandt and Otto, 2005). ADPKD is the most frequent, occurring in 1 in 400 to –1 in 1000 live births and affecting about 5 million persons worldwide (Torres et al., 2007). Other CDKs such as autosomal recessive polycystic kidney disease, pleiotropic Bardet–Biedl syndrome, Alstrom syndrome, and Orofaciodigital syndrome type 1, are by contrast, very infrequent diseases. CDKs are among the most common genetic lethal diseases has made them one of the most interesting and most studied subjects in epithelial biology. Despite the decades of long effort, the disease mechanisms remain mostly unknown. Polycystic kidney cells are characterized by increased proliferation and apoptosis of collector duct epithelial cells and a renal concentrating defect in spite of aquaporin 2 and vasopressin V2 receptor expression. Positional cloning helped to identify the first two CDK genes, PKD1 and PKD2 that carried mutations in ADPKD type 1 or 2, respectively (Mochizuki et al., 1996). Localization studies proved that PKD1 and PKD2 proteins, polycystin-1 and polycystin-2, are widely expressed in epithelia, vascular smooth muscle, cardiac myocytes and other tissues, and are regulated through development. Both proteins are mainly present in the primary cilia in wildtype cells (Yoder et al., 2002). In particular, polycystin-1 is located at the plasma membrane and especially at the primary cilium, whereas polycystin2 is located mainly at the primary cilium and the endoplasmic reticulum but also at the basolateral plasma membrane. Cilia are membrane protrusions sustained over a tubular structure conformed by nine microtubule doublets, and originate from the basal bodies, a similar microtubular structure that is attached to the cellular centrosome. Nonmotile (nonflagellar) primary cilia are modular sensory organelles that allow vertebrate cells to receive chemical and physical stimuli of diverse nature. During cloning of other CDK genes and cystic phenotype related genes, it was found that their products localized at the primary cilium, basal bodies or centrosomes, or were involved in their biogenesis (Watnick and Germino, 2003). It has been proposed that both proteins form a complex that functions as a sensor for mechanic stress (Nauli et al., 2003). Polycystin-1 and polycystin-2 are large membrane glycoproteins (Hughes et al., 1995; Ward et al., 1996), with 11 and 6 transmembrane domains, respectively. Polycystin-1 binds other membrane proteins through its extracellular domains and polycystin-2 works as a calcium channel. When a mechanic stimulus, such as

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urine flow, pushes the primary cilium, protein interactions change polycystin-1 conformation and polycystin-2 increases calcium permeability. Increases in intracellular calcium effect as a second messenger and produce a series of changes in cell physiology (Clapham, 2007; Nowycky and Thomas, 2002). Mutations impairing the ciliary pathway would deplete cellular Ca2þ levels affecting the subsequent cascade of mechanisms (Rizzuto and Pozzan, 2003). Ca2þ levels regulate inversin recruitment at the centrosome (Simons et al., 2005). When calcium levels increase, inversin is released and targets Dishevelled (Dvl) for degradation, thus switching off Wnt signaling, a cellular pathway that promotes cell cycle through upregulation of G1-phase genes. Ca2þ levels also affect cAMP levels by inhibiting the main renal adenylate cyclase, AC6 (Borodinsky and Spitzer, 2006; Cooper et al., 1995). Furthermore, calcium inhibits Ras activation through CAPRI RasGAP, and also inhibits cAMP-mediated MAPK signaling, a pathway that activates transcription of proliferation genes and Orofaciodigital syndrome type 1 (Cullen and Lockyer, 2002). Ca2þ depletion due to polycystin complex dysfunction would cause Wnt, Ras, and MAPK signaling activation and cell proliferation, an increase in cAMP levels and PKA-mediated induction of proapoptotic genes and finally a general defect in Ca2þ signaling. The polycystin complex is also part of focal adhesion complexes during epithelial cell migration in renal development and localizes at cell–cell junctions in fully developed renal tubules (Huan and van Adelsberg, 1999; Kreidberg et al., 1996; Roitbak et al., 2004). The mechanosensory hypothesis concludes that the complex could act as a sensor receiving mechanic signals from the matrix during migration, from partner cells during epithelial differentiation, and from the lumen of the developing organ via the primary cilium. Supporting this hypothesis, certain mutations in polycystin-1 induce focal adhesion kinase (FAK) mislocalization and defects in focal adhesion signaling, and developmental polycystin dysfunctions cause embryonic cystic kidneys, a severe phenotype with bad prognosis.

6. Concluding Remarks The establishment and maintenance of cell polarity is a critical step for the development of the epithelial identity. Cell polarity requires the communication of epithelial cells between them and with the surrounding tissues and ECM. Understanding the molecular mechanisms that regulates cell polarity is critical to comprehend normal tissue homeostasis, as well as the development and progression of malignant diseases. Alterations in epithelial tissues are the cause of diseases that can affect from just few individuals every year, such as the MID, to millions worldwide, such as

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cancer. In fact, cancer was the direct cause of almost 8 millions of deaths in 2007, which represent 1/8 of the total deaths worldwide. In the next future, work in in vivo models and in mammalian 3D culture model systems will provide us with a clearer understanding of the mechanisms of polarity regulation during epithelial morphogenesis, and how to apply this knowledge in medical treatments.

ACKNOWLEDGMENTS We thank Carmen Martin Ruiz-Jarabo for comments on the manuscript, and members of the Miguel A. Alonso and Isabel Correas lab for discussion. Work supported by grants to FM-B, from the Ministerio de Ciencia e Innovacio´n (BFU2008-01916), the European Union (MIRG-CT-2007-209382) and HFSP (LT00426/2004-C). AER-F is recipient of a MS fellowship from Fundacio´n Obra Social La Caixa. An institutional grant from the Fundacio´n Ramo´n Areces to CBMSO is also acknowledged.

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