Making the Connection: Ciliary Adhesion Complexes Anchor Basal Bodies to the Actin Cytoskeleton

Developmental Cell

Article Making the Connection: Ciliary Adhesion Complexes Anchor Basal Bodies to the Actin Cytoskeleton Ioanna Antoniades,1,2 Panayiota Stylianou,1,2 and Paris A. Skourides1,* 1Department

of Biological Sciences, University of Cyprus, P.O. Box 20537, Nicosia 2109, Cyprus authors contributed equally to this work *Correspondence: [email protected] http://dx.doi.org/10.1016/j.devcel.2013.12.003 2These

SUMMARY

Cilia have been associated with diverse developmental and physiological processes, and defects in cilia underlie a number of genetic conditions. Several lines of evidence support a critical role of the actin cytoskeleton in ciliogenesis and ciliary function. Here, we show that well-characterized focal adhesion (FA) proteins, including FAK, Paxillin, and Vinculin, associate with the basal bodies of multiciliated cells and form complexes (CAs) that interact with the actin cytoskeleton. FAK downregulation leads to ciliogenesis defects similar to those observed when the actin cytoskeleton is disrupted, including defects in basal body migration, docking, and spacing, suggesting that CAs link basal bodies to the actin cytoskeleton. The important role of FA proteins in ciliogenesis leads us to propose that evolutionarily FA proteins, many of which are found in primitive flagellated unicellular eukaryotes, may have originally evolved to perform functions at flagella and were later co-opted for use in cell adhesion. INTRODUCTION Cilia are microtubule-based organelles extending from basal bodies on the surface of vertebrate cells. There are two main types of cilia; primary and motile cilia. While primary cilia can be found on almost every cell type, motile cilia are restricted to specialized epithelia and are essential for generating fluid flow. Such epithelia include the airways, the ventricles of the brain, and the oviducts (Knowles and Boucher, 2002; Sawamoto et al., 2006; Shah et al., 2009; Worthington and Cathcart, 1963). Motile cilia can be found in single copy, as motile monocilia, or multicilia (Goetz and Anderson, 2010; Pazour and Witman, 2003; Pedersen et al., 2008; Roy, 2009), and the disruption of ciliary function has been shown to be responsible for a variety of human diseases termed ciliopathies (Afzelius, 1976, 2004; Baker and Beales, 2009; Bisgrove and Yost, 2006; Hildebrandt et al., 2011; Wallingford and Mitchell, 2011; Zariwala et al., 2007). Perhaps the most striking results regarding the importance of cilia come from the fly, where DSas-4 mutant larvae, which lack centrioles, develop into morphologically

normal adults, which die shortly after birth because their sensory neurons lack cilia (Basto et al., 2006). The dependence of ciliogenesis on the actin cytoskeleton is well documented, and both components of the actin cytoskeleton, as well as regulators of actin dynamics, have been shown to play important roles in various aspects of ciliogenesis (Bershteyn et al., 2010; Boisvieux-Ulrich et al., 1990; Dawe et al., 2007; Ioannou et al., 2013; Klotz et al., 1986; Lemullois et al., 1988; Pan et al., 2007; Ravanelli and Klingensmith, 2011; Tamm and Tamm, 1988). In multiciliated cells, basal bodies form de novo deep within the cytoplasm (Sorokin, 1968) and are subsequently transported, via an actin-myosin-based mechanism, to the apical surface where they dock (Boisvieux-Ulrich et al., 1990; Dawe et al., 2007; Klotz et al., 1986; Lemullois et al., 1987). In addition to the proposed role of the actin cytoskeleton in the transport of basal bodies, the apical surface of multiciliated cells is enriched with a dense meshwork of actin composed of two distinct pools, the apical and the subapical actin networks. Loss of the apical actin network leads to problems with basal body localization and polarity and appears to be necessary for basal body docking (Boisvieux-Ulrich et al., 1990; Park et al., 2006, 2008; Werner et al., 2011). Once basal bodies dock at the apical surface, they must be polarized in order for the cilia to beat in a synchronized manner and create directional fluid flow. The subapical actin network has been shown to be important for basal body spacing and metachronal synchrony. This network appears to connect basal bodies through the striated rootlets. Interestingly, disruption of the subapical actin network during basal body docking results in severe spacing issues, suggesting that spacing is determined by this actin network, while its disruption after ciliogenesis is completed leads to loss of metachornal synchrony (Werner et al., 2011). The actin cytoskeleton and its regulators have also been shown to influence the formation of primary cilia. Although the precise role of actin on primary cilia formation is not clear, studies suggest that modulation of the actin cytoskeleton regulates basal body positioning and primary cilium development overall, suggesting that primary cilia formation is promoted through the disassembly of certain pools of highly dynamic filamentous actin (Bershteyn et al., 2010; Kim et al., 2010; Sharma et al., 2011; Yan and Zhu, 2013; Yin et al., 2009). However, a mutation in Talpid3 has been shown to elicit defects in primary cilia formation and actin organization in the chicken. In these mutants, docking of basal bodies to the apical cell membrane is defective, despite the fact that mature basal bodies form (Yin et al., 2009). These

70 Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc.

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 1. FA Proteins FAK, Paxillin, and Vinculin Associate with Basal Bodies in Ciliated Cells (A and A0 ) Multiciliated cell expressing GFP FAK and centrin2 RFP. Each basal body is associated with a GFP FAK punctum at its posterior (in relation to the tadpole’s anterior-posterior axis). GFP FAK signal displays a gradient within the cell (in respect to the anterior-posterior axis), with elevated signal at the basal bodies localized at the cell’s posterior. (B and B0 ) Multiciliated cell expressing GFP Paxillin and centrin2 RFP. GFP Paxillin exhibits a similar localization and distribution throughout the cell, like GFP FAK. (C and C0 ) Multiciliated cell expressing GFP Vinculin and centrin2 RFP. Localization and distribution of GFP Vinculin resembles that of GFP FAK and GFP Paxillin. (D) Immunofluorescence staining with a p-S732 FAK antibody in centrin2 CFP-injected embryos. Endogenous FAK, associated with the basal bodies, is phosphorylated on Serine 732. (E) Immunofluorescence staining of endogenous Vinculin in centrin2 RFP-expressing multiciliated cells, gives a similar localization as GFP Vinculin. (F and F0 ) GRP ciliated cells of a stage 17 embryo expressing GFP FAK and centrin2 RFP, immunostained for acetylated tubulin. (G and G0 ) Neural tube cross section of a stage 24 embryo coexpressing GFP FAK and centrin2 RFP and immunostained for acetylated tubulin. GFP FAK associates with the basal bodies in both GRP and neural tube ciliated cells. (H and I) Immunofluorescence staining of endogenous Serine 732 phosphorylated FAK (H) and Paxillin (I) in NIH 3T3 cells, costained for g tubulin and acetylated tubulin. Endogenous FAK and Paxillin localize at the base of primary cilia next to the basal body in mammalian ciliated cells. See also Figure S1.

results suggest that formation of the apical actin network affects the attachment of basal bodies to the cell membrane and that certain aspects of the role of the actin network may be shared between motile and primary cilia. In addition to the studies in animals, actin has also been shown to be important for the control of flagellar length in Chlamydomonas, while knockdown of actin in Giardia results in defects in the organism’s flagella and their positioning, suggesting that the requirement of actin for normal ciliogenesis extends to flagellated unicellular eukaryotes (Dentler and Adams, 1992; Paredez et al., 2011). Despite the critical role of the actin cytoskeleton in ciliogenesis and ciliary function, how the basal bodies interact with the actin cytoskeleton is not known. Here, we show that four well-characterized focal adhesion (FA) proteins are found associated with basal bodies and form what we termed ‘‘ciliary adhesions’’ (CAs), connecting basal bodies to the actin cytoskeleton of multiciliated cells. Similar complexes are also found in monociliated cells of the gastrocel roof plate (GRP) and primary cilia. CA protein FAK interacts with both basal bodies and actin, while its downregulation leads to defects in ciliogenesis related to actin-based processes, suggesting that CAs functionally link basal bodies to the actin cytoskeleton.

RESULTS FAK, Paxillin, Vinculin, and Talin Are Associated with the Basal Bodies in Ciliated Cells FAs are large, dynamic protein complexes through which the actin cytoskeleton connects to the extracellular matrix. FAK and Paxillin are well-characterized FA proteins with critical roles in the assembly and disassembly of FA complexes. The fact that both FAK and Paxillin have been previously detected in association with centrosomes raised the possibility that FA proteins also associate with basal bodies (Herreros et al., 2000; Park et al., 2009). Using green fluorescent protein (GFP) fusions of FAK, Paxillin, and Vinculin in combination with live imaging, we noted that all three proteins localize in close association with the basal bodies, visualized using centrin2 red fluorescent protein (RFP), in Xenopus multiciliated cells (Figures 1A–1C0 ). Control cells expressing GFP alone show that GFP by itself does not localize at the basal bodies (Figures S1A and S1A0 available online), and western blotting showed that expression levels of GFP Paxillin and GFP Vinculin were similar to those of the endogenous proteins (Figures S1B and S1C). In addition, a fourth FA protein, Talin, fused to FusionRed also localizes at the basal bodies of multiciliated cells (Figure S1D). To further verify the GFP fusion

Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc. 71

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 2. FA Proteins Present an AnteriorPosterior Gradient in Multiciliated Cells (A–B) Intensity color-coded maximum intensity projections of centrin2 RFP (A) and GFP Vinculin (A0 ) reveals the presence of a Vinculin gradient in multiciliated cells. The signal of centrin2 RFP is uniform (A), while the GFP Vinculin signal is stronger at the cell’s posterior (A0 ). In (B), multiciliated cells coexpressing GFP Vinculin and centrin2 RFP are shown. The anterior-posterior gradient is maintained at the tissue level. The arrows indicate cells’ posterior where the signal for GFP Vinculin appears elevated. (C) Multiciliated cell coexpressing mKate2 FAK, clamp GFP, and centrin2 CFP. FAK is localized next to the basal body at the end of the region marked by clamp GFP. (D and E) Confocal optical sections of a multiciliated cell showing that GFP Paxillin and mKate2 FAK colocalize. GFP Paxillin is more concentrated at the apical-most region of the basal bodies (D), while mKate2 FAK is more concentrated slightly below (E). (F–G0 ) Intensity coding of ROIs of (D) and (E). Paxillin exhibits highest intensity at 0.00 mm, as shown in (F) and (F0 ), whereas FAK exhibits highest intensity at 0.38 mm, as shown in (G) and (G0 ).

localizations, we went on to stain endogenous FAK and Vinculin using previously characterized antibodies against these two proteins. Since FAK associated with centrosomes was shown to be phosphorylated on Serine 732, we used a phospho-Serine 732-specific FAK antibody in tadpoles expressing centrin2 cyan fluorescent protein (CFP), and, as shown, the staining confirms that endogenous FAK is associated with basal bodies and is phosphorylated on Serine 732 (Figure 1D). In addition, immunofluorescence experiments using a monoclonal Vinculin antibody in centrin2 RFP-expressing tadpoles gives very similar results to the GFP fusion (Figure 1E). We went on to express GFP FAK in neural tissues to examine the possibility that FA proteins also associate with basal bodies in the monociliated cells of the GRP and the primary cilia of the neural tube. As shown in Figures 1F and 1G, GFP FAK localizes adjacent to the basal bodies of both motile monocilia of the GRP (Figures 1F and 1F0 ) and primary cilia of the neural tube (Figures 1G and 1G0 ). The localization of FAK and Paxillin fusion constructs at the basal bodies of primary cilia was confirmed using antibodies to image the endogenous proteins in NIH 3T3 cells after serum-starvation-induced ciliogenesis (Figures 1H and 1I). These results suggest that the aforementioned FA proteins associate with the basal bodies in all types of cilia. FAK, Paxillin, and Vinculin Display Polarity within Multiciliated Cells of the Xenopus Epidermis In addition to the strong localization at the basal bodies, FAK, Paxillin, and Vinculin display polarity within the cell. Two types of polarity have been identified in multiciliated cells, both controlled by the planar cell polarity pathway: rotational polarity and tissue-level polarity (Mitchell et al., 2007, 2009; Park et al., 2008; Wallingford, 2010). Rotational polarity refers to the alignment of basal bodies within each cell, and the aforementioned FA proteins are clearly rotationally polarized (Figures 1A–1C).

Tissue-level polarity, on the other hand, refers to the coordination of the multiciliated cells across the tissue. However, FAK, Paxillin, and Vinculin also display a third type of polarity, with stronger signal emanating from basal bodies at the cell posterior and weaker signal from basal bodies at the front of the cell (in relation to the tadpole’s anterior-to-posterior axis) (Figures 1A–1C). This gradient is sharper in the case of Vinculin, in which the first row of basal bodies, in close proximity to the cortical actin cytoskeleton, shows a much higher level of Vinculin compared to the second row, which is not interacting with the cell cortex (Figures 1C and 1C0 ). This gradient becomes clearer in images where both centrin as well as Vinculin are presented using intensity coding, revealing a strong gradient in the cell (Figures 2A and 2A0 ). Imaging ciliated cells at lower magnifications shows that this polarity is retained at the tissue level (Figure 2B). The signal for all four proteins localizes immediately adjacent to each basal body and is rotationally polarized, but its relation to accessory structures, like the basal foot and the rootlet, cannot be determined without additional markers. To better define this region, we expressed mKate2 FAK with clamp GFP, to mark the striated rootlets, and centrin2 CFP, to mark the basal bodies. As shown in Figure 2C, the FAK signal is concentrated at the end of the region marked by clamp GFP, roughly, the area where the basal foot forms and an area where electron microscopy analysis identified interactions between basal bodies and the actin cytoskeleton (Chailley et al., 1989). This shows that the gradient within each cell described earlier, is an anterior-to-posterior gradient with higher concentrations at the basal bodies at the back of each cell (in relation to the tadpole’s anterior-to-posterior axis). Despite the fact that the localization of the three proteins was nearly identical, we noticed that Paxillin and Vinculin were always appearing simultaneously with the basal bodies in z stacks, while FAK appeared immediately after. This suggests that Paxillin and

72 Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc.

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 3. FA Proteins Form CA Complexes Connecting Basal Bodies to the Actin Network (A) Basal bodies (labeled with centrin2 RFP) are docked at the apical surface of multiciliated cells, in the same plane as apical actin (visualized with GFP utrophin). Each basal body is found at the center of an actin ring. (B) mKate2 FAK is in contact with the apical actin network. (C) Coexpression of centrin2 RFP and GFP FAK followed by phalloidin staining suggests that FAK connects basal bodies to the apical actin network. (D and D0 ) Multiciliated cell expressing GFP FAK, marking CAs, and mKate2 actin, showing the apical actin network, before (D) and after (D0 ) acceptor photobleaching. (D00 ) mKate2 (acceptor) intensity drops after photobleaching, while GFP (donor) intensity rises, showing that FRET is taking place between GFP and mKate2 and suggesting that FAK is interacting with actin. (E and E0 ) Multiciliated cell expressing centrin GFP and mKate2 actin before (E) and after (E0 ) acceptor photobleaching. (E00 ) mKate2 intensity drops after photobleaching, but the GFP intensity remains unchanged, suggesting that no FRET is taking place between GFP and mKate2, indicating the absence of a direct interaction between centrin and actin. See also Figure S2.

Vinculin display different distributions along the apicobasal axis compared to FAK. In an effort to confirm this, we coexpressed GFP Paxillin, mKate2 FAK, and centrin2 yellow fluorescent protein (YFP). As shown in Figures 2D and 2E, FAK and Paxillin colocalize but have a slightly different distribution along the z axis. Paxillin appears to have the highest density at the apical-most region of the basal bodies (Figures 2F and 2F0 ), while FAK has a maximal density slightly below the apical-most region of each basal body (Figures 2G and 2G0 ), as shown in the two adjacent optical sections of Figures 2F–2G0 . CAs Link Basal Bodies to the Actin Cytoskeleton The presence of four FA proteins in close association with basal bodies in a region shown to have interactions with the actin cytoskeleton raised the possibility that these proteins are forming a complex connecting basal bodies to the actin network. As shown in Figure 3A, each basal body is in the center of a ring of actin. On the other hand, FAK partially colocalizes with the apical actin network, as shown in Figure 3B, suggesting that FAK, Paxillin, Talin, and Vinculin are forming a complex linking each basal body to the actin network. Coexpression of GFP FAK with centrin2 RFP and staining with phalloidin shows that FAK is in contact with both the basal bodies and the actin network (Figure 3C). In order to confirm an interaction between FAK and the actin network, we coexpressed GFP FAK (donor) with mKate2 actin (acceptor) and carried out acceptor photobleaching experiments to examine the possibility of intermolecular fluorescence resonance energy transfer (FRET). These experiments show that FRET is taking place between GFP and mKate2 in ciliated cells (Figures 3D–3D00 ). Control experiments using centrin GFP as the donor and mKate2

actin as an acceptor fail to detect FRET, as expected, and show that centrin does not interact with actin, in agreement with the localization data (Figures 3E–3E00 ). Spot bleaching experiments confirm that FRET is restricted to the areas containing basal bodies (Figures S2A–S2A0 and Figure S2B). These results provide evidence that FA proteins form complexes that we termed CAs and that CAs are connecting each basal body to the actin network. Given the proposed role of actin in basal body transport and the close association of basal bodies and actin during this process, we wanted to explore the possibility that CAs are present during basal body migration (Boisvieux-Ulrich et al., 1990; Ioannou et al., 2013). Imaging basal bodies (centrin2 RFP) during ciliated cell intercalation revealed that CAs (marked by GFP Paxillin) are present during basal body migration, suggesting that CAs are likely connecting basal bodies to the internal actin network thought to be responsible for basal body transport (Figures 4A–4A000 and 4B) (Ioannou et al., 2013). Since basal bodies also connect to the subapical actin network through the ciliary rootlets, we went on to examine whether FA proteins are also involved in making this connection. As shown in Figure 4C0 , we found a second complex forming below the basal bodies, in close association with the subapical actin, suggesting that CAs are responsible for the connection between the ciliary rootlets and the subapical actin network. Coexpression of GFP FAK with clamp RFP and centrin2 CFP confirms that FAK is found both in association with the basal bodies, at the apical surface, as well as at the end of the ciliary rootlets (Figures 4D and 4D0 ). In order to get a better resolution of the association of the CAs with the actin cytoskeleton, both apical and subapical,

Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc. 73

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 4. CA Complexes Link Basal Bodies and Ciliary Rootlets to the Subapical Actin Network (A–A00 0 ) Optical sections of an intercalating multiciliated cell expressing GFP Paxillin and centrin2 RFP showing that Paxillin is associated with basal bodies during their migration to the apical surface. (B) 3D reconstruction (y-z) of optical sections of the intercalating cell in (A)–(A00 0 ). (C and C0 ) Multiciliated cell expressing GFP FAK and stained with phalloidin. GFP FAK exhibits the highest density at two focal planes. The first one corresponds to the plane of the apical actin network (C), while the second one appears slightly below, at the plane of the subapical actin network (C0 ). GFP FAK is in close association with both pools of actin. (D) Maximum intensity projection of a region of a multiciliated cell coexpressing centrin2 RFP (to label basal bodies), clamp RFP (to label the striated rootlet), and GFP FAK. GFP FAK exhibits two intensity maxima: one at the level of the cell’s apical surface, adjacent to the basal bodies, and a second one deeper, at the end of the ciliary rootlets. (D0 ) Confocal image from a mechanically sectioned (along the apicobasal axis) multiciliated cell coexpressing centrin2 CFP, clamp RFP, and GFP FAK. The arrows mark the two maxima of GFP FAK, one associated with the basal body and one associated with the end of the striated rootlet. (E) Optical section of a multiciliated cell coexpressing centrin2 CFP and mKate2 FAK and labeled with phalloidin (apical surface is at the top). The cell was initially mechanically sectioned, approximately along its anterior-posterior axis. The subapical actin, known to connect each basal body with the end of the striated rootlet of the cilium behind (Werner et al., 2011), appears to project from the CAs (red-labeled with mKate2 FAK) and form the characteristic discontinuous network. (F) Optical section of a multiciliated cell expressing mKate2 FAK and labeled with phalloidin (apical surface is at the top). The cell was initially mechanically sectioned, approximately along its left-to-right axis. The subapical actin appears to connect CAs of neighboring basal bodies (along the left-right axis), creating continuous loops. Again, the subapical actin appears to project from the CAs. (G) Multiciliated cell coexpressing GFP FRNK and centrin2 RFP. GFP FRNK associates with the basal bodies in a similar way as the full-length protein. It is localized at the posterior site of each basal body and exhibits an anterior-posterior gradient within the cell. (H) Multiciliated cell coexpressing Paxillin C GFP and centrin2 RFP. Paxillin C GFP localizes at the posterior site of basal bodies and displays a gradient with respect to the anterior-posterior axis of the cell.

we sectioned embryos expressing mKate2 FAK and stained them with phalloidin. This led to an improvement of the resolution in the apicobasal axis, since lateral resolution is higher than axial in confocal systems. We observed that the subapical network in these cells is projecting from the CAs basally and toward the back of the cell. We also observed that the subapical network, as described elsewhere (Werner et al., 2011), is connecting each basal body to the one immediately behind and in front through the rootlet, creating the characteristic discontinuous actin network shown in Figure 4E. However, we also noted an additional subapical network, rotated 90 with respect to the first, creating loops between the CAs of adjacent basal bodies (Figure 4F). This network appears to connect basal bodies in the left-to-right direction and is probably responsible for the spacing of basal bodies in this direction, while the discontinuous front-to-back subapical network is responsible for the spacing of basal bodies front to back. Both networks appear to be originating at apical CAs and are linked subapically at a nexus, presumably the subapical region showing elevated FAK concentrations at the end of the clamp-positive domain.

FAK is known to localize to FAs through its C-terminal region and, specifically, the FAT domain. This localization is believed to be strongly dependent on FAK’s interaction with Paxillin, which maps at the C terminus of FAK (Hayashi et al., 2002; Hildebrand et al., 1993, 1995). We went on to explore the domains required for FAK localization at the basal bodies. As shown in Figure 4G, the FAK C terminus, including the two proline-rich sequences and the FAT domain, (FRNK: FAK-Related NonKinase) is sufficient for the basal body localization of the protein. Similarly, the C terminus of Paxillin is also sufficient to drive FA localization of the protein (Brown et al., 1996) and, as shown in Figure 4H, is also sufficient for basal body localization, showing that, for both proteins, the determinants for CA complex localization are similar to those for FA localization, further emphasizing the similarity between the two complexes. Morpholino-Based FAK Downregulation Blocks Ciliogenesis Although the localization and FRET data are strongly suggestive of the function of the CA complex, we wanted to determine what effects the disruption of the complex would have on

74 Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc.

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 5. FAK Knockdown Leads to Defects in Ciliogenesis (A–C) Surface view (maximum intensity projection) of the epidermis of stage 31/32 Xenopus tadpoles stained with phalloidin and immunostained for acetylated tubulin, which stain actin network and cilia, respectively. In (A), cells of control embryos appear normal, with a lot of cilia projecting from their apical surface. In (B), morphant cells (of embryos injected at one ventral blastomere, at stage 5, with 4 ng of FAK MO) have fewer cilia projecting outward from their surface. In (C), coinjection of HA FAK mRNA (90 pg) with the FAK MO (4 ng) rescues the phenotype and cells project cilia normally. (D) Surface view (maximum intensity projection) of the epidermis of a stage 31/32 Xenopus embryo stained with phalloidin and immunostained for acetylated tubulin. Coinjection of HA-FAK D375 (45 pg) mRNA with the FAK MO does not rescue the ciliogenesis defects. (E) Quantification of the FAK MO induced ciliogenesis defects. Bar chart shows the percentage of stage 31/32 embryos that present mild, moderate, and severe defects or are normal with respect to the number of cilia projecting toward their surface. Data were collected from 130–150 cells, from ten embryos (three experiments) of each category (control, MO injected, MO and HA FAK injected, and MO and HA FAK D375 injected). (F–I) 3D reconstruction (x-z) of optical sections of ciliated cells expressing centrin2 RFP and clamp GFP. In (F), most basal bodies have reached the apical surface but are not docked properly in cells injected with 4 ng of FAK MO. In (G), higher amounts of FAK MO (8 ng) lead to a more dramatic phenotype, as the majority of basal bodies fail to reach the apical surface of the cell. In (H), all the basal bodies are docked at the apical surface of a control ciliated cell. In (I), coinjection of HA FAK mRNA (120 pg) with the FAK MO (8 ng) rescues the phenotype, as the basal bodies have managed to reach the apical surface and the majority of them have docked properly. (J and K) Intercalating ciliated cells of stage 17 Xenopus embryos coexpressing centrin2 RFP and GFP utrophin to label basal bodies and the actin network, respectively. In control cells (J), basal bodies are clustered deep in the cell, surrounded by an actin network, and individual basal bodies associate with actin filaments. In cells coinjected with 8 ng of FAK MO (K), the basal bodies are dispersed, and many of them do not associate with actin, which fails to form an organized network around them.

ciliogenesis. Since FAK is a well-characterized regulator of the dynamics of FA assembly and disassembly, we postulated that it would have an important role in CA complex function. We thus used a previously characterized FAK morpholino (MO) oligo (Fonar et al., 2011; Petridou et al., 2012, 2013) (after verifying its efficiency at tadpole stages; Figure S3A) to block FAK expression in multiciliated cells. As shown in Figure 5B, FAK morphant cells have few cilia projecting from their surface compared to control cells (Figures 5A and 5E). Coinjection of FAK mRNA with the FAK MO effectively rescues this phenotype, and rescued cells project cilia normally, suggesting that

the phenotype is specific (Figures 5C and 5E). Since the FERM domain of FAK is critical for its regulation and has recently been shown to have an important role in the regulation of FAK’s dynamics and function at FAs, we decided to examine if the FERM domain was required for rescue (Cooper et al., 2003; Petridou et al., 2013). We thus coinjected the FAK D375 mutant, which lacks the FERM domain, with the FAK MO. The FAK D375 mutant failed to rescue the phenotype, showing that the FERM domain and/or proper regulation of FAK’s kinase activity are required for FAK’s function at CAs (Figures 5D and 5E).

Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc. 75

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 6. Apical Actin Enrichment and Rotational Polarity Are Unaffected in FAK Morphants (A–C) Confocal images of phalloidin-stained control (A), morphant (B), and rescued (C) ciliated cells expressing centrin2 RFP plus enlarged close-ups of selected regions, shown in (A0 ), (B0 ), and (C0 ), and a respective deeper optical section of each, in (A00 ), (B00 ), and (C00 ), showing details of apical and subapical actin networks. Morphants (B), unlike controls (A) and rescued (C) cells, display reduced numbers of basal bodies at the cell surface with large areas completely devoid of basal bodies. The apical actin network is present both in morphants and controls; however, it appears less organized and less dense in regions devoid of basal bodies. The subapical actin of morphants (B00 ) is completely missing in areas devoid of basal bodies and appears disorganized in areas with basal bodies. (D and E) Centrin2 RFP- and clamp GFP-expressing control (D) and FAK morphant (E) multiciliated cells. Spacing is disrupted, with regions of the apical surface devoid of basal bodies in the morphant cell. However, overall rotational polarity is not significantly affected as most rootlets are oriented in the same direction. See also Figure S3.

Examination of centrin in side projections of morphant cells (Figure 5F) reveals that, despite the fact that most basal bodies reach the apical surface, they display variations in positioning in the z axis, unlike the controls (Figure 5H), suggesting that many of them do not dock. At higher doses of the FAK MO (8 ng), basal body transport is impaired and the majority of basal bodies remain deep in the cytosol (Figure 5G). Imaging high-dose morphant cells during intercalation reveals that basal bodies are often dispersed, rather than clustered like in controls, and some clearly fail to associate with the internal actin network, previously implicated in their transport (BoisvieuxUlrich et al., 1990; Ioannou et al., 2013), suggesting that the transport defect is due to loss of association between the basal bodies and the actin network (Figures 5J and 5K). Coinjection of FAK mRNA also rescues the transport phenotype as shown in Figure 5I. In addition to the ciliogenesis and docking defects, FAK morphants (4 ng) display spacing issues with regions of the apical surface completely devoid of basal bodies (Figure 6B). Despite the presence of apical actin enrichment in these regions, the network appears less dense (Figure 6B0 ) and the subapical actin is completely lost (Figure 6B00 ), unlike control (Figures 6A–6A00 ) and rescued cells (Figures 6C–6C00 ), which present uniformly organized apical and subapical actin networks. In addition, although the apical actin of morphant cells appears normal at regions with docked basal bodies, the subapical actin appears less organized. Interestingly, the use of clamp GFP with centrin2 RFP shows that rotational polarity, which is

dependent on the microtubule network, remains relatively unaffected in FAK morphants, especially when examining local coordination in basal body clusters (Figures 6D and 6E). These results suggest that FAK has an important role in the regulation of the CA complex, and in addition, they are consistent with a role of this complex as a mechanical link between basal bodies and the actin cytoskeleton. Since the C terminus of FAK (FRNK) is sufficient for CA localization, we postulated that, in a similar fashion as in FAs, it would be able to act as a dominant negative by displacing endogenous FAK (Gilmore and Romer, 1996; Hildebrand et al., 1993; Schaller et al., 1992; Taylor et al., 2001). We went on to overexpress FRNK in order to further address the role of FAK and CAs during ciliogenesis. Expression of FRNK partially displaces FAK from CAs but fails to induce strong ciliogenesis defects (Figures S3B and S3C; data not shown). In FRNK-expressing cells, most basal bodies reach the apical surface and project cilia; however, in these cells, basal body spacing is disrupted (Figures S3D–S3F; data not shown). Since the subapical actin network has been shown to be responsible for basal body spacing, this result supports a role of the CA complex linking basal bodies to the actin network (Werner et al., 2011). The reduced severity of the FRNK phenotype compared to the FAK MO can be explained by the fact that FRNK fails to displace all endogenous FAK from CAs. In addition, FRNK only blocks kinase-dependent functions of FAK while retaining scaffolding functions that depend on the C-terminal FAT domain and the two proline-rich regions of the protein (Gilmore and Romer, 1996; Hildebrand et al., 1993; Taylor et al., 2001). This result is in agreement with a role of CA complexes as a mechanical link between actin and

76 Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc.

Developmental Cell CAs Link Basal Bodies to the Actin Network

Figure 7. CA Complexes in Multiciliated Cells (A) A diagram showing the localization of the CA complexes with respect to the basal bodies, ciliary rootlets, and the actin cytoskeleton in multiciliated cells. CA complexes interact with the apical and subapical pools of actin. The upper red circle shows an x-z projection of confocal optical sections of CA complexes and basal bodies. CAs are marked with GFP FAK (green) and the basal bodies with centrin2 RFP (red). (B) Proposed arrangement of the subapical actin network in multiciliated cells in relation to the basal bodies, rootlets, and CAs.

the basal bodies and, in addition, suggests that FAK has important kinase-independent roles at the CAs. DISCUSSION Overall, this work reveals a function of FA proteins in the creation of complexes responsible for anchoring basal bodies to the actin network of multiciliated cells. In addition, our results show that CAs also form at the ends of ciliary rootlets and are responsible for connecting the subapical actin network to the ciliary rootlet. We also show that the subapical network has two components. One discontinuous network in the frontto-back direction connects each basal body to the one in front and the one behind through the ciliary rootlet; and a continuous network, at a right angle in relation to the first, connects adjacent basal bodies in the left-to-right direction. This network is probably responsible for spacing of basal bodies in the left-to-right orientation while the discontinuous front-to-back

network determines spacing in the anterior-posterior axis of the cell. The location of CAs with respect to the basal bodies, rootlets, and the actin network is summarized in Figure 7. The localization of the apical CA complex at the cell’s posterior makes the most sense mechanically, since the maximum force would be excreted by the cilium on the basal bodies during the effective stroke rather than the recovery stroke, pushing the basal bodies in the opposite direction of the stroke. Linking the basal body to the actin network on this area would ensure the immobilization of the basal body during the effective stroke. The similarities between CAs and FAs are striking, and the determinants for Paxillin and FAK localization on CAs appear to be similar to those for FA localization, although a more detailed analysis will be required to determine if they are, in fact, identical. FAK has been shown to be a critical regulator of FA assembly and disassembly. Loss of FAK activity leads to aberrant ciliogenesis and defects that are dependent on actin-based processes, like basal body transport, docking, and spacing, without affecting apical actin enrichment. These results suggest that FAK, a critical regulator of FAs, is also an important regulator of CAs, and they are consistent with a role of this complex in the mechanical linkage of basal bodies to the actin cytoskeleton. The proximity of the apical CAs to the basal foot also raises the possibility that CAs are also connecting basal bodies to the microtubule network, shown to be responsible for establishing the rotational polarity within multiciliated cells. However, FAK morphants show no rotational polarity defects and, in this respect, resemble cytochalasin-treated cells where spacing is affected but local rotational polarity is maintained, making it unlikely that CAs are linking basal bodies to microtubules (Werner et al., 2011). The presence of CA complexes in both motile and primary cilia suggests that these proteins serve a highly conserved function. The role of the actin cytoskeleton is well documented both for motile cilia and primary cilia, as well as the flagella of unicellular eukaryotes (Dentler and Adams, 1992; Engel et al., 2011; Kim et al., 2010; Paredez et al., 2011; Sharma et al., 2011). Interestingly, several core FA proteins have been found in flagellated unicellular eukaryotes, including Apusozoa and Choanoflagellates, the closest sister group of Metazoa (Sebe´-Pedro´s et al., 2010). Flagellar motility is found in every major eukaryotic group (Fritz-Laylin et al., 2010) and is believed to be an ancestral feature present in the last common ancestor of all eukaryotic organisms (Carvalho-Santos et al., 2011; Fritz-Laylin et al., 2010; Mitchell, 2007). This raises the possibility that the core FA proteins originally evolved to perform functions at flagella and were later co-opted for use in cell adhesion. Further support for this notion comes from the fact that FA proteins were lost in all fungi, with the exception of chytrid fungi, which are the only members of the kingdom with flagellated gametes, suggesting an important function for FA proteins in flagellar function, in agreement with our results (Carvalho-Santos et al., 2011). Future work will focus on examining the possibility that CAs also exist in flagellated unicellular eukaryotes and will also focus on addressing the precise role of individual proteins within this complex.

Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc. 77

Developmental Cell CAs Link Basal Bodies to the Actin Network

EXPERIMENTAL PROCEDURES Embryo Manipulations and Microinjections Female adult Xenopus laevis were ovulated by injection of human chorionic gonadotropin. Eggs were fertilized in vitro, dejellied in 2% cysteine (pH 7.8), and subsequently reared in 0.1 3 Marc’s modified ringers (MMR) and staged according to Neiuwkoop and Faber (Nieuwkoop and Faber, 1994). For microinjections, embryos were placed in a solution of 4% Ficoll in 0.33 3 MMR and injected using a glass capillary pulled needle, forceps, a Singer Instruments MK1 micromanipulator, and a Harvard Apparatus pressure injector. After injections, embryos were reared for 2 hr or until stage 8 in 4% Ficoll in 0.33 3 MMR and then washed and maintained in 0.1 3 MMR. For most experiments, injections were made into the ventral blastomeres to target the epidermis at the 4-cell, 8-cell, or 16-cell stage. For all experiments, we injected morpholinos at 4 to 8 ng per blastomere and mRNAs at various amounts. Embryos were allowed to develop to the appropriate stage and then imaged live or fixed in MEMFA (Sive et al., 2010) for 1–2 hr at room temperature (RT). Fixed embryos were used immediately. For live imaging, embryos were anesthetized in 0.01% benzocaine in 0.1 3 MMR. Immunostaining Indirect immunofluorescence assays were carried out as described elsewhere (Demetriou et al., 2008; Skourides et al., 1999) with modifications. NIH 3T3 cells were seeded on glass coverslips (charged with HCl), washed three times with PBS, preextracted with 0.2% Triton X-100 for 20 s, and then fixed for 10 min in 4% paraformaldehyde solution in PBS. Fixation was followed by addition of 50 mM glycine solution in PBS, and then the cells were blocked using 10% normal goat serum (Jackson Immunoresearch) for 30 min. Cells were incubated with primary antibodies diluted in 10% normal goat serum solution in PBS for 1.5 hr. The primary antibodies used were Paxillin mouse monoclonal (BD Transduction Laboratories) used in combination with Acetyl-a-Tubulin (Lys40) rabbit polyclonal (Cell Signaling) and g Tubulin goat polyclonal (Santa Cruz Biotechnology); and FAK [pS732] Phosphospecific rabbit polyclonal (Invitrogen) used in combination with Acetylated tubulin, mouse monoclonal (Santa Cruz Biotechnology) and g Tubulin goat polyclonal (Santa Cruz Biotechnology). Cells were then washed five times in PBS. Secondary antibodies used were Cy3 anti-mouse and anti-rabbit (Jackson Immunoresearch), Alexa 488 anti-goat (Molecular Probes Invitrogen), and IgG-CFL 647 anti-mouse and anti-rabbit (Santa Cruz Biotechnology). For whole-mount immunostaining, fixed embryos were permeabilized in PBDT (1 3 PBS + 0.5% Triton X-100 + 1% DMSO) for several hours at RT and blocked in PBDT + 1% normal goat serum for 1 hr at RT. Primary antibodies were added in block solution, and embryos were incubated for 4 hr at RT or overnight at 4 C. The primary antibodies used were: Acetylated tubulin mouse monoclonal (Santa Cruz Biotechnology), Vinculin mouse monoclonal (Developmental Studies Hybridoma Bank), and FAK pS732 rabbit polyclonal (Invitrogen). The next day, embryos were washed 4 3 10 min in PBDT. Embryos were then incubated in secondary antibodies—anti-mouse Alexa 488 (Invitrogen) or anti-mouse IgG-CFL 647 (Santa Cruz Biotechnology)—at RT for 1–2 hr. Actin labeling was performed using Alexa Fluor Phalloidin 488 (Invitrogen) and Phalloidin CruzFluor 647 (Santa Cruz Biotechnology) at RT for 1–2 hr. Embryos were then washed 4 3 10 min in PBDT. Embryos were postfixed in MEMFA for 10-15 min at RT, washed in 1 3 PBS and imaged immediately. Western Blot Analysis Protein lysates were prepared by homogenizing embryos (by pipetting up and down) in ice-cold MK’s modified lysis buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 0.5% NP-40, 0.5% Triton X-100, 100 mM EGTA, 5 mM NaF) supplemented with protease inhibitors (1 mM phenylmethylsulfonyl fluoride and protease cocktail; Sigma). Homogenates were cleared by centrifugation at 15,000 3 g for 30 min at 4 C (Kragtorp and Miller, 2006). Proteins (3/4–1 embryo equivalent) were run on 7.5% SDS-polyacrylamide gels with the WesternC ladder (Bio-Rad) and blotted on to nitrocellulose membranes. Blots were blocked in 5% milk (in TBSTw: 13 TBS buffer and 0.1% Tween 20) and then incubated with the primary antibodies in 5% milk overnight at 4 C. The primary antibodies used were: FAK mouse monoclonal (Millipore), Paxillin

mouse monoclonal (BD Transduction Laboratories), and Vinculin mouse monoclonal (Developmental Studies Hybridoma Bank). Visualization was performed using horseradish peroxidase-conjugated antibodies (anti-rabbit and anti-mouse; Santa Cruz Biotechnology), after 1 hr incubation at RT, and signal was detected using LumiSensor (GeneScript) on a UVP BioSpectrum Imaging System. For loading control, actin rabbit polyclonal antibody (Santa Cruz Biotechnologies) was used. GRP Assay Embryos were fixed in MEMFA at stage 17. The GRP was manually dissected and postfixed for a further 15 min. GRP tissue was then used for immunofluorescence as described earlier. Imaging and Quantification Imaging was performed on a Zeiss LSM 710 laser scanning confocal microscope with Zen 2010 software. Quantification of the ciliogenesis defects was conducted based on the number of cilia projecting from the apical surface of each cell. Cells have been denoted as normal, with mild phenotype (having more than 50% of their cilia but less than control cells), with moderate phenotype (having 50% or lower reduction of their cilia), and with severe phenotype (having near complete loss of their cilia). Quantification of the basal body distribution was conducted by overlaying a grid over each cell and counting the number of basal bodies included in each cell of the grid, using Adobe Photoshop CS2 software. Acceptor Photobleaching FRET FRET experiments were accomplished using a laser scanning confocal microscope (Zeiss LSM 710) with a Plan-Apochromat 633/1.40 Oil DIC M27 objective lens (Zeiss). Stage 33–35 embryos expressing GFP FAK and mKate2 actin or centrin GFP and mKate2 actin were anesthetized in 0.01% benzocaine in MMR and immobilized in silicone grease wells on glass slides. A 543 nm laser was used for acceptor (mKate2 actin) photobleaching (100% power) within a region of interest (ROI). A 488 nm laser (0.8% power) was used for acquisition of GFP FAK, and emission was detected between 493 and 538 nm. A 543 nm laser (1.2% power) was used for acquisition of mKate2 actin, and emission was detected between 599 and 758 nm. One frame was acquired as a prebleaching control, and the ROI (rectangular region) was bleached within one frame. Zeiss Zen 2010 software was used for FRET analysis. FRET efficiency was calculated using the following equation: FRETEfficiency = (Donor Post  Donor Pre)/(Donor Post) 3 100 (Donor Post: donor emission intensity after photobleaching; Donor Pre: donor emission intensity before photobleaching). SUPPLEMENTAL INFORMATION Supplemental Information includes Supplemental Experimental Procedures and three figures and can be found with this article online at http://dx.doi. org/10.1016/j.devcel.2013.12.003. ACKNOWLEDGMENTS We thank Drs. John Wallingford, Brian Mitchell, and Reinhard Ko¨ster for kindly providing plasmids. We also thank Dr. Niovi Santama for providing reagents for the immunofluorescence experiments on NIH 3T3 cells. Finally, we thank Charalambos Kourouklaris for generating the 3D reconstruction of the proposed model. Funding was provided by the Cyprus Research Promotion Foundation (YGEIA/BIOS/0609(BE)/14, TEXNOLOGIA/YLIKA/0311(BIE)/10). Accepted: December 6, 2013 Published: January 13, 2014 REFERENCES Afzelius, B.A. (1976). A human syndrome caused by immotile cilia. Science 193, 317–319. Afzelius, B.A. (2004). Cilia-related diseases. J. Pathol. 204, 470–477.

78 Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc.

Developmental Cell CAs Link Basal Bodies to the Actin Network

Baker, K., and Beales, P.L. (2009). Making sense of cilia in disease: the human ciliopathies. Am. J. Med. Genet. C. Semin. Med. Genet. 151C, 281–295. Basto, R., Lau, J., Vinogradova, T., Gardiol, A., Woods, C.G., Khodjakov, A., and Raff, J.W. (2006). Flies without centrioles. Cell 125, 1375–1386. Bershteyn, M., Atwood, S.X., Woo, W.M., Li, M., and Oro, A.E. (2010). MIM and cortactin antagonism regulates ciliogenesis and hedgehog signaling. Dev. Cell 19, 270–283.

Hildebrand, J.D., Schaller, M.D., and Parsons, J.T. (1995). Paxillin, a tyrosine phosphorylated focal adhesion-associated protein binds to the carboxyl terminal domain of focal adhesion kinase. Mol. Biol. Cell 6, 637–647. Hildebrandt, F., Benzing, T., and Katsanis, N. (2011). Ciliopathies. N. Engl. J. Med. 364, 1533–1543.

Bisgrove, B.W., and Yost, H.J. (2006). The roles of cilia in developmental disorders and disease. Development 133, 4131–4143.

Ioannou, A., Santama, N., and Skourides, P.A. (2013). Xenopus laevis nucleotide binding protein 1 (xNubp1) is important for convergent extension movements and controls ciliogenesis via regulation of the actin cytoskeleton. Dev. Biol. 380, 243–258.

Boisvieux-Ulrich, E., Laine´, M.C., and Sandoz, D. (1990). Cytochalasin D inhibits basal body migration and ciliary elongation in quail oviduct epithelium. Cell Tissue Res. 259, 443–454.

Kim, J., Lee, J.E., Heynen-Genel, S., Suyama, E., Ono, K., Lee, K., Ideker, T., Aza-Blanc, P., and Gleeson, J.G. (2010). Functional genomic screen for modulators of ciliogenesis and cilium length. Nature 464, 1048–1051.

Brown, M.C., Perrotta, J.A., and Turner, C.E. (1996). Identification of LIM3 as the principal determinant of paxillin focal adhesion localization and characterization of a novel motif on paxillin directing vinculin and focal adhesion kinase binding. J. Cell Biol. 135, 1109–1123.

Klotz, C., Bordes, N., Laine, M.C., Sandoz, D., and Bornens, M. (1986). Myosin at the apical pole of ciliated epithelial cells as revealed by a monoclonal antibody. J. Cell Biol. 103, 613–619.

Carvalho-Santos, Z., Azimzadeh, J., Pereira-Leal, J.B., and Bettencourt-Dias, M. (2011). Evolution: Tracing the origins of centrioles, cilia, and flagella. J. Cell Biol. 194, 165–175.

Knowles, M.R., and Boucher, R.C. (2002). Mucus clearance as a primary innate defense mechanism for mammalian airways. J. Clin. Invest. 109, 571–577.

Chailley, B., Nicolas, G., and Laine´, M.C. (1989). Organization of actin microfilaments in the apical border of oviduct ciliated cells. Biol. Cell 67, 81–90.

Kragtorp, K.A., and Miller, J.R. (2006). Regulation of somitogenesis by Ena/VASP proteins and FAK during Xenopus development. Development 133, 685–695.

Cooper, L.A., Shen, T.L., and Guan, J.L. (2003). Regulation of focal adhesion kinase by its amino-terminal domain through an autoinhibitory interaction. Mol. Cell. Biol. 23, 8030–8041.

Lemullois, M., Klotz, C., and Sandoz, D. (1987). Immunocytochemical localization of myosin during ciliogenesis of quail oviduct. Eur. J. Cell Biol. 43, 429–437.

Dawe, H.R., Farr, H., and Gull, K. (2007). Centriole/basal body morphogenesis and migration during ciliogenesis in animal cells. J. Cell Sci. 120, 7–15.

Lemullois, M., Boisvieux-Ulrich, E., Laine, M.C., Chailley, B., and Sandoz, D. (1988). Development and functions of the cytoskeleton during ciliogenesis in metazoa. Biol. Cell 63, 195–208.

Demetriou, M.C., Stylianou, P., Andreou, M., Yiannikouri, O., Tsaprailis, G., Cress, A.E., and Skourides, P. (2008). Spatially and temporally regulated alpha6 integrin cleavage during Xenopus laevis development. Biochem. Biophys. Res. Commun. 366, 779–785. Dentler, W.L., and Adams, C. (1992). Flagellar microtubule dynamics in Chlamydomonas: cytochalasin D induces periods of microtubule shortening and elongation; and colchicine induces disassembly of the distal, but not proximal, half of the flagellum. J. Cell Biol. 117, 1289–1298. Engel, B.D., Ishikawa, H., Feldman, J.L., Wilson, C.W., Chuang, P.T., Snedecor, J., Williams, J., Sun, Z., and Marshall, W.F. (2011). A cell-based screen for inhibitors of flagella-driven motility in Chlamydomonas reveals a novel modulator of ciliary length and retrograde actin flow. Cytoskeleton (Hoboken) 68, 188–203. Fonar, Y., Gutkovich, Y.E., Root, H., Malyarova, A., Aamar, E., Golubovskaya, V.M., Elias, S., Elkouby, Y.M., and Frank, D. (2011). Focal adhesion kinase protein regulates Wnt3a gene expression to control cell fate specification in the developing neural plate. Mol. Biol. Cell 22, 2409–2421. Fritz-Laylin, L.K., Prochnik, S.E., Ginger, M.L., Dacks, J.B., Carpenter, M.L., Field, M.C., Kuo, A., Paredez, A., Chapman, J., Pham, J., et al. (2010). The genome of Naegleria gruberi illuminates early eukaryotic versatility. Cell 140, 631–642. Gilmore, A.P., and Romer, L.H. (1996). Inhibition of focal adhesion kinase (FAK) signaling in focal adhesions decreases cell motility and proliferation. Mol. Biol. Cell 7, 1209–1224. Goetz, S.C., and Anderson, K.V. (2010). The primary cilium: a signalling centre during vertebrate development. Nat. Rev. Genet. 11, 331–344. Hayashi, I., Vuori, K., and Liddington, R.C. (2002). The focal adhesion targeting (FAT) region of focal adhesion kinase is a four-helix bundle that binds paxillin. Nat. Struct. Biol. 9, 101–106. Herreros, L., Rodrı´guez-Fernandez, J.L., Brown, M.C., Alonso-Lebrero, J.L., Caban˜as, C., Sa´nchez-Madrid, F., Longo, N., Turner, C.E., and Sa´nchezMateos, P. (2000). Paxillin localizes to the lymphocyte microtubule organizing center and associates with the microtubule cytoskeleton. J. Biol. Chem. 275, 26436–26440. Hildebrand, J.D., Schaller, M.D., and Parsons, J.T. (1993). Identification of sequences required for the efficient localization of the focal adhesion kinase, pp125FAK, to cellular focal adhesions. J. Cell Biol. 123, 993–1005.

Mitchell, D.R. (2007). The evolution of eukaryotic cilia and flagella as motile and sensory organelles. Adv. Exp. Med. Biol. 607, 130–140. Mitchell, B., Jacobs, R., Li, J., Chien, S., and Kintner, C. (2007). A positive feedback mechanism governs the polarity and motion of motile cilia. Nature 447, 97–101. Mitchell, B., Stubbs, J.L., Huisman, F., Taborek, P., Yu, C., and Kintner, C. (2009). The PCP pathway instructs the planar orientation of ciliated cells in the Xenopus larval skin. Curr. Biol. 19, 924–929. Nieuwkoop, P.D.F., and Faber, J. (1994). Normal Table of Xenopus laevis (Daudin): A Systematical and Chronological Survey of the Development from the Fertilized Egg till the End of Metamorphosis, First Edition. (New York: Garland). Pan, J., You, Y., Huang, T., and Brody, S.L. (2007). RhoA-mediated apical actin enrichment is required for ciliogenesis and promoted by Foxj1. J. Cell Sci. 120, 1868–1876. Paredez, A.R., Assaf, Z.J., Sept, D., Timofejeva, L., Dawson, S.C., Wang, C.J., and Cande, W.Z. (2011). An actin cytoskeleton with evolutionarily conserved functions in the absence of canonical actin-binding proteins. Proc. Natl. Acad. Sci. USA 108, 6151–6156. Park, T.J., Haigo, S.L., and Wallingford, J.B. (2006). Ciliogenesis defects in embryos lacking inturned or fuzzy function are associated with failure of planar cell polarity and Hedgehog signaling. Nat. Genet. 38, 303–311. Park, T.J., Mitchell, B.J., Abitua, P.B., Kintner, C., and Wallingford, J.B. (2008). Dishevelled controls apical docking and planar polarization of basal bodies in ciliated epithelial cells. Nat. Genet. 40, 871–879. Park, A.Y., Shen, T.L., Chien, S., and Guan, J.L. (2009). Role of focal adhesion kinase Ser-732 phosphorylation in centrosome function during mitosis. J. Biol. Chem. 284, 9418–9425. Pazour, G.J., and Witman, G.B. (2003). The vertebrate primary cilium is a sensory organelle. Curr. Opin. Cell Biol. 15, 105–110. Pedersen, L.B., Veland, I.R., Schrøder, J.M., and Christensen, S.T. (2008). Assembly of primary cilia. Dev. Dyn. 237, 1993–2006. Petridou, N.I., Stylianou, P., Christodoulou, N., Rhoads, D., Guan, J.L., and Skourides, P.A. (2012). Activation of endogenous FAK via expression of its amino terminal domain in Xenopus embryos. PLoS ONE 7, e42577.

Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc. 79

Developmental Cell CAs Link Basal Bodies to the Actin Network

Petridou, N.I., Stylianou, P., and Skourides, P.A. (2013). A dominant-negative provides new insights into FAK regulation and function in early embryonic morphogenesis. Development 140, 4266–4276. Ravanelli, A.M., and Klingensmith, J. (2011). The actin nucleator Cordon-bleu is required for development of motile cilia in zebrafish. Dev. Biol. 350, 101–111. Roy, S. (2009). The motile cilium in development and disease: emerging new insights. Bioessays 31, 694–699. Sawamoto, K., Wichterle, H., Gonzalez-Perez, O., Cholfin, J.A., Yamada, M., Spassky, N., Murcia, N.S., Garcia-Verdugo, J.M., Marin, O., Rubenstein, J.L., et al. (2006). New neurons follow the flow of cerebrospinal fluid in the adult brain. Science 311, 629–632.

Sorokin, S.P. (1968). Reconstructions of centriole formation and ciliogenesis in mammalian lungs. J. Cell Sci. 3, 207–230. Tamm, S., and Tamm, S.L. (1988). Development of macrociliary cells in Beroe¨. I. Actin bundles and centriole migration. J. Cell Sci. 89, 67–80. Taylor, J.M., Mack, C.P., Nolan, K., Regan, C.P., Owens, G.K., and Parsons, J.T. (2001). Selective expression of an endogenous inhibitor of FAK regulates proliferation and migration of vascular smooth muscle cells. Mol. Cell. Biol. 21, 1565–1572. Wallingford, J.B. (2010). Planar cell polarity signaling, cilia and polarized ciliary beating. Curr. Opin. Cell Biol. 22, 597–604.

Schaller, M.D., Borgman, C.A., Cobb, B.S., Vines, R.R., Reynolds, A.B., and Parsons, J.T. (1992). pp125FAK a structurally distinctive protein-tyrosine kinase associated with focal adhesions. Proc. Natl. Acad. Sci. USA 89, 5192–5196.

Wallingford, J.B., and Mitchell, B. (2011). Strange as it may seem: the many links between Wnt signaling, planar cell polarity, and cilia. Genes Dev. 25, 201–213.

Sebe´-Pedro´s, A., Roger, A.J., Lang, F.B., King, N., and Ruiz-Trillo, I. (2010). Ancient origin of the integrin-mediated adhesion and signaling machinery. Proc. Natl. Acad. Sci. USA 107, 10142–10147.

Werner, M.E., Hwang, P., Huisman, F., Taborek, P., Yu, C.C., and Mitchell, B.J. (2011). Actin and microtubules drive differential aspects of planar cell polarity in multiciliated cells. J. Cell Biol. 195, 19–26.

Shah, A.S., Ben-Shahar, Y., Moninger, T.O., Kline, J.N., and Welsh, M.J. (2009). Motile cilia of human airway epithelia are chemosensory. Science 325, 1131–1134.

Worthington, W.C., Jr., and Cathcart, R.S., 3rd. (1963). Ependymal cilia: distribution and activity in the adult human brain. Science 139, 221–222.

Sharma, N., Kosan, Z.A., Stallworth, J.E., Berbari, N.F., and Yoder, B.K. (2011). Soluble levels of cytosolic tubulin regulate ciliary length control. Mol. Biol. Cell 22, 806–816. Sive, H.L., Grainger, R.M., and Richard, M.H. (2010). Early Development of Xenopus laevis: A Laboratory Manual. (Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press). Skourides, P.A., Perera, S.A., and Ren, R. (1999). Polarized distribution of Bcr-Abl in migrating myeloid cells and co-localization of Bcr-Abl and its target proteins. Oncogene 18, 1165–1176.

Yan, X., and Zhu, X. (2013). Branched F-actin as a negative regulator of cilia formation. Exp. Cell Res. 319, 147–151. Yin, Y., Bangs, F., Paton, I.R., Prescott, A., James, J., Davey, M.G., Whitley, P., Genikhovich, G., Technau, U., Burt, D.W., and Tickle, C. (2009). The Talpid3 gene (KIAA0586) encodes a centrosomal protein that is essential for primary cilia formation. Development 136, 655–664. Zariwala, M.A., Knowles, M.R., and Omran, H. (2007). Genetic defects in ciliary structure and function. Annu. Rev. Physiol. 69, 423–450.

80 Developmental Cell 28, 70–80, January 13, 2014 ª2014 Elsevier Inc.