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• W1799771027 abstract "Article18 May 2012free access Identification of a novel Wnt5a–CK1ε–Dvl2–Plk1-mediated primary cilia disassembly pathway Kyung Ho Lee Kyung Ho Lee Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USAPresent address: Chemical Biology Research Center, Korea Research Institute of Bioscience and Biotechnology, Republic of Korea Search for more papers by this author Yoshikazu Johmura Yoshikazu Johmura Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA Search for more papers by this author Li-Rong Yu Li-Rong Yu Division of Systems Biology, Center for Proteomics, National Center for Toxicological Research, US Food and Drug Administration, Jefferson, AR, USA Search for more papers by this author Jung-Eun Park Jung-Eun Park Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA Search for more papers by this author Yuan Gao Yuan Gao Division of Systems Biology, Center for Proteomics, National Center for Toxicological Research, US Food and Drug Administration, Jefferson, AR, USA Search for more papers by this author Jeong K Bang Jeong K Bang Division of Magnetic Resonance, Korea Basic Science Institute, Chung-Buk, Republic of Korea Search for more papers by this author Ming Zhou Ming Zhou Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., Frederick National Laboratory for Cancer Research, Frederick, MD, USA Search for more papers by this author Timothy D Veenstra Timothy D Veenstra Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., Frederick National Laboratory for Cancer Research, Frederick, MD, USA Search for more papers by this author Bo Yeon Kim Bo Yeon Kim Chemical Biology Research Center and World Class Institute, Korea Research Institute of Bioscience and Biotechnology, Chung-Buk, Republic of Korea Search for more papers by this author Kyung S Lee Corresponding Author Kyung S Lee Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA Search for more papers by this author Kyung Ho Lee Kyung Ho Lee Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USAPresent address: Chemical Biology Research Center, Korea Research Institute of Bioscience and Biotechnology, Republic of Korea Search for more papers by this author Yoshikazu Johmura Yoshikazu Johmura Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA Search for more papers by this author Li-Rong Yu Li-Rong Yu Division of Systems Biology, Center for Proteomics, National Center for Toxicological Research, US Food and Drug Administration, Jefferson, AR, USA Search for more papers by this author Jung-Eun Park Jung-Eun Park Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA Search for more papers by this author Yuan Gao Yuan Gao Division of Systems Biology, Center for Proteomics, National Center for Toxicological Research, US Food and Drug Administration, Jefferson, AR, USA Search for more papers by this author Jeong K Bang Jeong K Bang Division of Magnetic Resonance, Korea Basic Science Institute, Chung-Buk, Republic of Korea Search for more papers by this author Ming Zhou Ming Zhou Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., Frederick National Laboratory for Cancer Research, Frederick, MD, USA Search for more papers by this author Timothy D Veenstra Timothy D Veenstra Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., Frederick National Laboratory for Cancer Research, Frederick, MD, USA Search for more papers by this author Bo Yeon Kim Bo Yeon Kim Chemical Biology Research Center and World Class Institute, Korea Research Institute of Bioscience and Biotechnology, Chung-Buk, Republic of Korea Search for more papers by this author Kyung S Lee Corresponding Author Kyung S Lee Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA Search for more papers by this author Author Information Kyung Ho Lee1, Yoshikazu Johmura1, Li-Rong Yu2, Jung-Eun Park1, Yuan Gao2, Jeong K Bang3, Ming Zhou4, Timothy D Veenstra4, Bo Yeon Kim5 and Kyung S Lee 1 1Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, Bethesda, MD, USA 2Division of Systems Biology, Center for Proteomics, National Center for Toxicological Research, US Food and Drug Administration, Jefferson, AR, USA 3Division of Magnetic Resonance, Korea Basic Science Institute, Chung-Buk, Republic of Korea 4Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., Frederick National Laboratory for Cancer Research, Frederick, MD, USA 5Chemical Biology Research Center and World Class Institute, Korea Research Institute of Bioscience and Biotechnology, Chung-Buk, Republic of Korea *Corresponding author. Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute/NIH, 9000 Rockville Pike, Building 37, Room 3118, Bethesda, MD 20892-4258, USA. Tel.:+1 301 496 9635; Fax:+1 301 496 8419; E-mail: [email protected] The EMBO Journal (2012)31:3104-3117https://doi.org/10.1038/emboj.2012.144 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Non-motile primary cilium is an antenna-like structure whose defect is associated with a wide range of pathologies, including developmental disorders and cancer. Although mechanisms regulating cilia assembly have been extensively studied, how cilia disassembly is regulated remains poorly understood. Here, we report unexpected roles of Dishevelled 2 (Dvl2) and interphase polo-like kinase 1 (Plk1) in primary cilia disassembly. We demonstrated that Dvl2 is phosphorylated at S143 and T224 in a manner that requires both non-canonical Wnt5a ligand and casein kinase 1 epsilon (CK1ε), and that this event is critical to interact with Plk1 in early stages of the cell cycle. The resulting Dvl2–Plk1 complex mediated Wnt5a–CK1ε–Dvl2-dependent primary cilia disassembly by stabilizing the HEF1 scaffold and activating its associated Aurora-A (AurA), a kinase crucially required for primary cilia disassembly. Thus, via the formation of the Dvl2–Plk1 complex, Plk1 plays an unanticipated role in primary cilia disassembly by linking Wnt5a-induced biochemical steps to HEF1/AurA-dependent cilia disassembly. This study may provide new insights into the mechanism underlying ciliary disassembly processes and various cilia-related disorders. Introduction Primary cilia, observed in most stromal and epithelial cells, are microtubule-based cellular sensing structures important for transducing extracellular signals. Aberrant ciliary function leads to many ciliopathies, such as renal cysts, hypertension, diabetes, neuronal, visual, respiratory, and other developmental disorders (Davenport and Yoder, 2005; Fliegauf et al, 2007; Adams et al, 2008; Sharma et al, 2008; Berbari et al, 2009). Unlike motile cilia, primary cilia lack a pair of central microtubules and dynein arms (9+0 arrangements of microtubules), and thus do not show wave motion. Primary cilia undergo a dynamic cycle of assembly and disassembly throughout the cell cycle. They assemble from basal body/mother centriole in G0- or G1-phase and rapidly disassemble prior to mitosis entry (Dawe et al, 2007; Pan and Snell, 2007; Santos and Reiter, 2008; Seeley and Nachury, 2010). Over the years, various components important for primary cilia assembly have been isolated and mechanisms underlying this event have begun to emerge. However, how the ciliary disassembly process is regulated remains poorly understood. Given that cancer cells commonly lack primary cilia (Michaud and Yoder, 2006; Plotnikova et al, 2008), pathways regulating cilia assembly/disassembly processes could be tightly linked to the mechanisms regulating tumourigenesis. Polo-like kinase 1 (Plk1) plays a pivotal role during the M-phase of the cell cycle (Barr et al, 2004; van de Weerdt and Medema, 2006; Petronczki et al, 2008; Archambault and Glover, 2009). Plk1 specifically localizes to centrosomes, kinetochores, and midbody through the function of the C-terminal non-catalytic polo-box domain (PBD) (Park et al, 2010), which forms a conserved phospho-Ser/Thr (pS/pT)-binding module (Cheng et al, 2003; Elia et al, 2003). Plk1 is highly overexpressed in various human cancers and is thought to promote tumourigenesis (Strebhardt and Ullrich, 2006). Although Plk1 does not appear to be required for proper ciliogenesis (Soung et al, 2009), whether it contributes to the ciliary disassembly process is not known. The Wnt signalling pathway is involved in a wide range of developmental processes, such as cell migration, proliferation, and polarity. Deregulation of Wnt signalling is associated with various human diseases, including cancer (Logan and Nusse, 2004; Moon et al, 2004; Clevers, 2006). Binding of various Wnt ligands to Frizzled (Fz), a family of G protein-coupled cell surface receptors, triggers signalling cascades, and Dishevelled (Dvl) plays a key role as a signal mediator in both canonical and non-canonical Wnt pathways (Moon et al, 2002; Nusse, 2005; Angers and Moon, 2009; Gao and Chen, 2010). Activation of the Wnt canonical pathway leads to stabilization of β-catenin, which turns on a set of genes by binding to LEF/TCF family transcriptional factors and regulates cell proliferation. Non-canonical pathways, which do not involve the function of β-catenin, also regulate various physiologically important processes, including planar cell polarity, axon guidance, and intracellular Ca++ release (Logan and Nusse, 2004; Zhang et al, 2007; Angers and Moon, 2009; Gao and Chen, 2010). Although molecular mechanisms are yet to be clarified, a flurry of evidence suggests that Wnt pathways are tightly associated with cilia assembly/disassembly processes (see reviews Gerdes and Katsanis, 2008; He, 2008). Casein kinase 1 (CK1) is a member of the Ser/Thr kinase superfamily with seven mammalian CK1 isoforms (α, β, γ1, γ2, γ3, δ, and ε) characterized to date (Knippschild et al, 2005). Among them, CK1ε exhibits a high enzyme-substrate specificity for Dvl phosphorylation in vitro and its closest relative CK1δ (82% identity in the entire amino-acid sequence and 96% identity in the kinase domain) also phosphorylates Dvl in vitro (McKay et al, 2001; Gao et al, 2002; Price, 2006; Dahlberg et al, 2009). Interestingly, both CK1δ and CK1ε appear to be required for Wnt5a-induced Dvl2 phosphorylation in vivo (Bryja et al, 2007). However, whether these phosphorylation events are physiologically significant, and whether CK1δ and CK1ε have distinct cellular functions remains largely elusive. In this study, we provide evidence that both non-mitotic Plk1 and Dvl2 have unanticipated roles in primary cilia disassembly. The PBD of Plk1 binds to the CK1ε-phosphorylated, conserved S143 and T224 sites of Dvl2, the most abundant isoform of the mammalian Dvl1–3 family (Lee et al, 2008), and this event is crucial in mediating the Wnt5a–CK1ε–Dvl2-dependent primary cilia disassembly pathway at the early stage of the cell cycle (note that S143 of Dvl2 is different from the previously identified S142 of Dvl1, which is analogous to S158 of Dvl2 (Klimowski et al, 2006)). Furthermore, the Dvl2–Plk1 complex appears to modulate HEF1 stability and, therefore, HEF1/Aurora-A (AurA)-dependent primary cilia disassembly, likely through the regulation of Smad3-dependent HEF1 degradation (Liu et al, 2000; Nourry et al, 2004). Given the close relationship between primary cilia disassembly and the cell cycle (Quarmby and Parker, 2005; Plotnikova et al, 2008; Seeley and Nachury, 2010) and the crucial role of Plk1 in promoting cell proliferation, we propose that Plk1 plays a key role in coupling primary cilia disassembly with cell-cycle progression. Results Identification of Dvl2 as a novel Plk1 PBD-binding protein To investigate new functions of Plk1 at the centrosome, we attempted to identify previously uncharacterized Plk1 PBD-binding proteins by performing PBD pull-downs with various centrosomal proteins expressed in asynchronously growing HeLa cells. Among the proteins that efficiently bind to the PBD was Dvl2, a component that plays a central role in mediating both canonical and non-canonical Wnt signalling pathways. Subsequent studies on the interaction between endogenous Plk1 and transfected Flag-tagged Dvl2 (Flag–Dvl2) showed that these two proteins were co-immunoprecipitated in a reciprocal manner (Figure 1A). Furthermore, GST–PBD efficiently interacted with both fast-migrating (i.e., underphosphorylated; marked ‘a’) and slow-migrating (i.e., hyperphosphorylated; marked ‘b’) Dvl2 forms in HeLa lysates prepared from asynchronously (Asy)-growing or thymidine (Thy)-treated (S-phase) cells (Figure 1B). The somewhat reduced level of Dvl2 bound to GST–PBD in nocodazole (Noc)-treated (M-phase) cells in Figure 1B appears to be due to inefficient detection of the hyperphosphorylated form of Dvl2, which was greatly improved by λ phosphatase treatment (Supplementary Figure S1). The respective GST–PBD (H538A K540M) mutant (PBD (AM)), defective in phospho-dependent binding (Elia et al, 2003), interacted with Dvl2 only marginally (Figure 1B). Consistent with this observation, provision of a previously characterized PBD-binding phosphopeptide, PLHSpT (Yun et al, 2009) or its high-affinity N-alkylphenyl derivatives, 4h and 4j (Liu et al, 2011), but not the non-phospho control PLHST peptide, disrupted the PBD–Dvl2 interaction (Figure 1C). Thus, Plk1 interacts with Dvl2 in a phospho-dependent manner throughout the cell cycle. Figure 1.Interaction between Plk1 PBD and the p-S143 and p-T224 epitopes of Dvl2. (A) HeLa cells transfected with Flag–Dvl2 were subjected to reciprocal immunoprecipitations. Immunoprecipitates were treated with λ phosphatase (PPase) to eliminate phosphorylated and slow-migrating forms and then immunoblotted. Asterisk, degradation product of Flag–Dvl2. (B) Total lysates prepared from asynchronously (Asy) growing, thymidine (Thy)-treated, or Noc-treated HeLa cells were subjected to pull-downs with bead-associated control GST, GST–PBD WT, or GST–PBD (H538A K540M) (AM). Precipitates were immunoblotted and the membrane was stained with Coomassie (CBB). Note that both fast- and slow-migrating Dvl2 forms (arbitrarily referred to as a and b forms, respectively) efficiently bound to GST–PBD (WT), but not to GST–PBD (AM). (C) Lysates from asynchronously growing HeLa cells were mixed with nonphospho-T78 peptide (PLHST), phospho-T78 peptide (PLHSpT), high-affinity PLHSpT derivatives 4h or 4j (Liu et al, 2011), or control buffer. The resulting lysates were then subjected to PBD pull-down as in (B). (D) Bead-immobilized Dvl2-derived peptides were incubated with HeLa lysates, precipitated, and then immunoblotted. (E) HEK293T cells transfected with the indicated Dvl2 constructs were subjected to PBD pull-downs, treated with λ phosphatase (PPase), and analysed. Numbers, relative amounts of Dvl2 bound to GST–PBD. (F) Anti-Flag–Dvl2 immunoprecipitates prepared from transfected HeLa cells arrested with thymidine for 18 h were subjected to mass spectrometry analysis to determine in-vivo phosphorylation site. The phosphorylation site was determined by the phosphorylated (red) MS/MS fragment ions with (w/) or without (w/o) neutral loss of phosphate and unphosphorylated (blue) fragment ions. The full results are shown in Supplementary Figure S2B. (G) HEK293T cells transfected with the indicated Dvl2 constructs were immunoblotted in the presence of 5 μg/ml of the indicated S143 or T224 peptide to determine the specificity of the phospho-antibodies. Figure source data can be found with the Supplementary data. Download figure Download PowerPoint Visual scanning of the Dvl2 primary sequence revealed multiple potential PBD-binding sequences that fall into the consensus PBD-binding motif (Φ/P-Φ-T/Q/H/M-S-pS/pT-P/X; X, any amino-acid residue; Φ, hydrophobic residue) (Elia et al, 2003). Testing of the phosphorylated peptides derived from these sequences showed that both p-S143 and p-T224 peptides efficiently interacted with Plk1, while p-T197 and p-S398 peptides interacted with Plk1 at a lesser level (Figure 1D). Among these four sites, mutation of S143 to A severely crippled the PBD–Dvl2 interaction, while mutation of T224 to A mildly weakened the interaction (Figure 1E). Mass spectrometry analyses with immunoprecipitated Flag–Dvl2 revealed that both S143 and T224 residues are phosphorylated in vivo (Figure 1F; Supplementary Figure S2A and B), which was further confirmed by immunoblotting analyses with phospho-specific antibodies generated against the two phospho-epitopes (Figure 1G). In line with the importance of S143 and T224 for PBD binding, both of these residues are highly conserved among vertebrates and also in different human isoforms (Supplementary Figure S2C and D). CK1δ and CK1ε induce the Dvl2–Plk1 interaction by phosphorylating the S143 and T224 residues on Dvl2 To determine the kinase(s) responsible for the phosphorylation of the S143 and T224 residues, HeLa cells were treated with previously characterized inhibitors. Among them, a CK1δ/ε-specific inhibitor, IC261 (Gao et al, 2002; Tillement et al, 2008), induced a fast-migrating Dvl2 form (a form) and greatly diminished the levels of both p-S143 and p-T224 epitopes (Figure 2A). In contrast, inhibition of GSK-3 (GSK inhibitor X), AurA and AurB (VX-680), Plk1 (BI 2536), or Cdc2 (BMI-1026) failed to significantly alter the levels of the p-S143 and p-T224 epitopes. Consistent with these findings, treatment of cells with IC261 abolished the Plk1 PBD–Dvl2 interaction (Figure 2B). Figure 2.CK1δ/ε-dependent phosphorylation of Dvl2 at S143 and T224. (A) Thymidine-arrested HeLa cells were treated with the indicated compounds to inhibit CK1δ/ε (IC261), GSK-3β (GSK Inhibitor X), AurA (VX-680), Plk1 (BI 2536), or Cdk (BMI-1026), respectively, and the resulting samples were subjected to immunoblotting analyses. Note only IC261 generated the fast-migrating ‘a’ form and greatly diminished the levels of the p-S143 and p-T224 epitopes. Numbers in (A) indicate relative signal intensities. (B) HeLa cells were treated with control DMSO or IC261 for 3 h and then subjected to PBD pull-downs. Precipitates were treated with λ phosphatase (PPase), immunoblotted, and the resulting membrane was stained with Coomassie (CBB). (C–E) Asynchronously growing hTERT-RPE cells were either silenced for control luciferase (GL), CK1δ, CK1ε, or both CK1δ and CK1ε by lentivirus-based shRNA (C, D) or transfected with CK1δ or CK1ε and treated with thymidine for 18 h (E). The resulting cells were then immunoblotted with the indicated antibodies (C, E) or subjected to PBD pull-downs (D). Where indicated, samples were separated by 15% SDS–PAGE to condense the Dvl2 signals for easy quantification or 8% low-bis-acrylamide gel to reveal hyperphosphorylated, slow-migrating proteins. Failure to induce an additive effect of shCK1δ and shCK1ε on the generation of the p-S143 and p-T224 epitopes (C) and in the PBD binding (D) could be in part attributable to the inefficient depletion of CK1ε (arrows). Numbers in (C) indicate signal intensities relative to the signal in the shGL sample. Note that the transfected CK1δ and CK1ε in (E) were overexpressed over their endogenous proteins ∼40 times and 10 times, respectively (data not shown). Figure source data can be found with the Supplementary data. Download figure Download PowerPoint Subsequent analyses showed that depletion of either CK1δ or CK1ε greatly diminished the levels of the p-S143 and p-T224 epitopes on Dvl2 (Figure 2C). However, significant levels of the p-S143 and p-T224 epitopes were still detectable in the cells depleted of both CK1δ and CK1ε (shCK1δ/ε). This could be due to the incomplete depletion of CK1δ or CK1ε (arrow) or to the presence of other unidentified kinase(s) that phosphorylates these sites. Notably, depletion of CK1δ induced the Dvl2, a form much more efficiently than depletion of CK1ε (Figure 2C; compare lane 2 with lane 3 in the 8% low-bis gel Dvl2 panel). This finding suggests that CK1δ and CK1ε-dependent Dvl2 phosphorylation sites do not completely overlap and that CK1δ and CK1ε may have distinct functions yet to be characterized. Consistent with the importance of CK1δ or CK1ε in the generation of the p-S143 and p-T224 epitopes, depletion of these kinases markedly impaired the PBD–Dvl2 binding (Figure 2D). In a second experiment, we observed that CK1δ and CK1ε directly phosphorylated and generated the p-S143 and p-T224 epitopes in vitro (Supplementary Figure S2E) and induced these phospho-epitopes in vivo (Figure 2E), thus corroborating the critical role of these kinases in the induction of the Dvl2–Plk1 complex. Plk1 promotes primary cilia disassembly in a kinase activity-dependent manner Since Dvl2 mediates both canonical and non-canonical Wnt pathways, we first investigated whether Plk1 contributes to the β-catenin-dependent canonical pathway. However, Plk1 activity neither altered the degree of β-catenin ubiquitination nor influenced the level of β-catenin-dependent transcription activity (Supplementary Figure S3), thereby diminishing the likelihood that Dvl2-bound Plk1 contributes to the canonical pathway. It has been suggested that Dvl2 regulates the apical docking and planar polarization of basal body in multiciliated epithelial cells and promotes ciliogenesis (Park et al, 2008). Thus, we investigated whether the formation of the Dvl2–Plk1 complex contributes to the processes of primary cilia assembly. Surprisingly, however, we observed that depletion of Dvl2 in hTERT-immortalized retinal pigment epithelial (hTERT-RPE) cells induced substantially longer primary cilia in most cells, whereas overexpression of Dvl2 induced markedly shorter cilia in a fewer cell population (Supplementary Figure S4A–E; see also Figure 4B–D, below). Similar results were obtained with NIH 3T3 mouse fibroblast cells (Supplementary Figure S4F–H). These findings suggest that Dvl2 facilitates the disassembly of primary cilia. In line with this view, depletion of Dvl2 in cells derived from a patient suffering from premature chromatid separation (PCS) syndrome also resulted in a longer primary cilia (Miyamoto et al, 2011). Therefore, we investigated whether Plk1 contributes to primary cilia disassembly and whether the formation of the Dvl2–Plk1 complex is important for this event. To this end, hTERT-RPE cells were infected with control shLuciferase (shGL) or shPlk1 lentivirus, and then serum starved for 48 h to allow the cells to generate primary cilia. The resulting cells were then stimulated with serum to induce primary cilia disassembly, harvested at various time points after stimulation, and then analysed. In the control shGL cells, the level of Plk1 became progressively abundant as the cells proceeded through the cell cycle, whereas the level of Plk1 remained low in the shPlk1 cells (Figure 3A). Under these conditions, the shGL cells gradually resorbed primary cilia over time. Twenty-four hours after serum stimulation, most (∼90%) of the population completely resorbed primary cilia, while the remaining population displayed much-shortened (∼2 μm) primary cilia (Figure 3B and C) (note that cells infected with lentivirus exhibited somewhat less efficient primary cilia formation in comparison to those in Figure 3D below; see also Supplementary Figure S4I). In stark contrast, only a small fraction (∼7%) of the shPlk1 population detectably resorbed primary cilia during the same period of time, and the rest displayed substantially longer (∼5 μm) cilia than the control shGL cells (Figure 3B and C; see also Figure 4D below). Consistent with a role of Plk1 in primary cilia disassembly, a significantly higher shPlk1 population exhibited primary cilia in Figure 3C. A separate experiment with much narrower time intervals suggested that Plk1 is required for proper primary cilia disassembly from as early as 4–6 h after serum stimulation in a sustained manner (Supplementary Figure S5A). Both of the shGL and shPlk1 cells entered the cell cycle efficiently following serum stimulation, as evidenced by the accumulated Cyclin A-positive population (Figure 3C) and by flow cytometry analyses (Supplementary Figure S5B; note that the shPlk1 cells do not show a detectable level of cell-cycle delay until ∼12–18 h after serum stimulation). These observations suggest that the observed delay in primary cilia disassembly in shPlk1 cells is not the consequence of an altered cell cycle in these cells. Measurement of Plk1 kinase activity in these cells revealed that the shGL cells, but not the shPlk1 cells, exhibited a gradually increasing level of Plk1 activity as the cells proceeded through the cell cycle (Figure 3C). Figure 3.Requirement of Plk1 activity for proper disassembly of primary cilia. (A, B) hTERT-RPE cells were infected with lentiviruses expressing shGL or shPlk1. Forty-eight hours after serum starvation, the cells were stimulated with serum, harvested, and then subjected to immunoblotting (A) and immunostaining (B) analyses. Magnified images of cilia are shown in the enlarged boxes in (B). (C) From the immunostained samples in (B), cells with primary cilia were quantified (bar graphs). A fraction of the total lysates in (A) was used to determine Plk1 activity using an ELISA-based assay (solid lines) (Park et al, 2009). To monitor cell-cycle progression, Cyclin A-positive cells were quantified (dotted lines). Error bars, standard deviation from more than three independent experiments. Note that a low but significant level of Plk1 activity was detectable even at the early stages after serum stimulation. (D) Control parental hTERT-RPE cells or hTERT-RPE Plk1-as cells lacking endogenous Plk1 but expressing the indicated constructs were starved for 48 h and stimulated with serum. The resulting cells were fixed at the indicated time points after stimulation and immunostained as in (B) to quantify the cells with primary cilia. Error bars, standard deviation. (E) Total lysates prepared from the 24-h samples in (D) were subjected to immunoblotting analyses and ELISA-based kinase assays (graph) to determine the level of Plk1 activity. Arrowhead indicates endogenous Plk1 that is not detectable in the hTERT-RPE Plk1-as cells deleted of the genomic PLK1 locus. Asterisks, EGFP–Plk1 degradation products. For graphs in (C, D), >300 cells were counted from the samples at each time point. Figure source data can be found with the Supplementary data. Download figure Download PowerPoint Figure 4.Induction of primary cilia disassembly by the CK1ε-dependent Plk1–Dvl2 p-S143/p-T224 complex. (A–D) hTERT-RPE cells infected with the indicated sh-lentiviruses were first starved for 48 h. A set of the resulting cells was harvested for immunoblotting (A) or immunostaining (B) analyses. Enlarged images of cilia are shown in the boxes at the bottom right corner (B). Another set of the starved cells was subsequently stimulated with serum, fixed at the indicated time point, and immunostained. The cells with primary cilia were then counted (C) and the average lengths of primary cilia among cilia-positive cells were quantified (D). Numbers in (A) indicate signal intensities relative to α-tubulin signals. Both anti-α-tubulin immunoblotting and Coomassie (CBB) staining were carried out for loading controls. Error bars, standard deviation from more than three independent experiments. Statistics: ***P<0.001 (unpaired two-tailed t-test). (E, F) hTERT-RPE cells expressing the indicated constructs were infected with lentivirus expressing either control shGL or shDvl2, and then immunoblotted (E). The resulting cells were starved for 48 h and then stimulated with serum. The cells harvested at the indicated time points after serum stimulation were immunostained and quantified (F). Note that cells infected with lentiviruses (expression or RNAi viruses) exhibited less efficient primary cilia formation. Asterisk, degradation product. Error bars, standard deviation from more than three independent experiments. For samples in (C, D, F), >300 cells were counted for each sample. Figure source data can be found with the Supplementary data. Download figure Download PowerPoint To directly determine whether Plk1 kinase activity is required for primary cilia disassembly, we expressed control vector, kinase-inactive Plk1 (K82M) (Lee et al, 1995), or Plk1 wild-type (WT) in the hTERT-RPE Plk1-as mutant cells lacking both copies of the PLK1 locus but expressing a greatly deactivated Plk1-as allele (Burkard et al, 2007) (see also Figure 3E, below). We observed that the Plk1-as mutant expressing either control vector or Plk1 (K82M) exhibited a considerably delayed primary cilia disassembly (Figure 3D). By contrast, Plk1-as cells expressing WT Plk1, which exhibited Plk1 kinase activity at a level similar to that of control hTERT-RPE WT cells, disassembled primary cilia as efficiently as the control WT cells (Figure 3D and E). It has been reported that a mitotic kinase, AurA, promotes primary cilia disassembly through the interaction with a pro-metastatic scaffolding protein, HEF1 (Pugacheva et al, 2007). Remarkably, treatment of cells with a Plk1 inhibitor, BI 2536, attenuated the ci" @default.
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• W1799771027 date "2012-05-18" @default.
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• W1799771027 title "Identification of a novel Wnt5a-CK1ε-Dvl2-Plk1-mediated primary cilia disassembly pathway" @default.
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