Advertisement
Short Report
Direct interaction between CEP85 and STIL mediates PLK4-driven directed cell migration
Yi Liu, Jaeyoun Kim, Reuben Philip, Vaishali Sridhar, Megha Chandrashekhar, Jason Moffat, Mark van Breugel, Laurence Pelletier
Journal of Cell Science 2020 133: jcs238352 doi: 10.1242/jcs.238352 Published 23 April 2020

ABSTRACT

PLK4 has emerged as a prime target for cancer therapeutics, and its overexpression is frequently observed in various types of human cancer. Recent studies have further revealed an unexpected oncogenic activity of PLK4 in regulating cancer cell migration and invasion. However, the molecular basis behind the role of PLK4 in these processes still remains only partly understood. Our previous work has demonstrated that an intact CEP85–STIL binding interface is necessary for robust PLK4 activation and centriole duplication. Here, we show that CEP85 and STIL are also required for directional cancer cell migration. Mutational and functional analyses reveal that the interactions between CEP85, STIL and PLK4 are essential for effective directional cell motility. Mechanistically, we show that PLK4 can drive the recruitment of CEP85 and STIL to the leading edge of cells to promote protrusive activity, and that downregulation of CEP85 and STIL leads to a reduction in ARP2 (also known as ACTR2) phosphorylation and reorganization of the actin cytoskeleton, which in turn impairs cell migration. Collectively, our studies provide molecular insight into the important role of the CEP85–STIL complex in modulating PLK4-driven cancer cell migration.

This article has an associated First Person interview with the first author of the paper.

INTRODUCTION

Cell migration is a fundamental cellular process essential for animals. In cancer, an unscheduled increase in the motile property of cancer cells can promote intravasation and metastasis, and this step-wise process relies on the spatiotemporal control of microtubule and actin dynamics (De Pascalis and Etienne-Manneville, 2017; Mayor and Etienne-Manneville, 2016; Paul et al., 2017). As the primary microtubule-organizing center, the centrosome plays a crucial role in the control of cell motility. During directional migration, centrosomes are repositioned towards the leading edge of cells to nucleate microtubules and to direct intracellular trafficking frontward (Ananthakrishnan and Ehrlicher, 2007; Rørth, 2007). Centrosomes also act as actin-organizing centers through pericentriolar material (PCM)-dependent recruitment of the WASP and Scar homolog (WASH) and ARP2/3 complex, providing an alternative mechanism for centrosome-based control of cell migration (Farina et al., 2016).

The centrosome consists of a pair of centrioles surrounded by PCM (Azimzadeh and Marshall, 2010; Gönczy, 2012). Centrioles must duplicate once per cell cycle to ensure that two centrosomes are present in mitosis to organize the spindle poles; failure to do so leads to a variety of diseases like cancer, microcephaly and ciliopathies (Strnad and Gönczy, 2008; Nigg and Raff, 2009; Arquint et al., 2014). PLK4 is one of the most upstream regulators of centriole duplication, and its kinase activity essential to initiate this process. Increased PLK4 activity leads to centriole amplification, while its inhibition prevents centriole formation (Habedanck et al., 2005; Kleylein-Sohn et al., 2007). PLK4 activity is tightly controlled through trans-autophosphorylation, which causes its degradation (Cunha-Ferreira et al., 2009; Guderian et al., 2010; Holland et al., 2010; Rogers et al., 2009). Additionally, direct interaction of STIL with PLK4 has been shown to further stimulate its kinase activity, possibly by promoting PLK4 autophosphorylation (Arquint and Nigg, 2016; Moyer et al., 2015; Ohta et al., 2014). More recently, we have reported that binding of CEP85 to STIL is critical for robust PLK4 activation and efficient centriole assembly (Liu et al., 2018).

PLK4-directed centrosome amplification is sufficient to induce aneuploidy and promote tumor initiation in flies and in mice when the p53 pathway is partially inhibited (Levine et al., 2017; Basto et al., 2008). Moreover, excess PLK4 expression has also been linked to a distinct cellular invasion mechanism through increased centrosomal-based microtubule nucleation and increase of RAC1 activity at the cortex (Godinho et al., 2014). A more recent study also revealed unexpected oncogenic activity for PLK4 in regulating ARP2/3-mediated actin rearrangement to promote cancer cell migration and invasion in a RAC1- and CDC42-independent manner (Kazazian et al., 2017). It still remains elusive, however, how PLK4 mediates contextual control of actin remodeling and cell motility. In addition, CEP192, an upstream regulator of PLK4, has been shown to participate in the control of directional migration (O'Rourke et al., 2014; Fung et al., 2018).

Here, we show that the CEP85–STIL protein module participates in directed cell motility. Using structure-guided information, we find that disrupting the CEP85–STIL binding interface negatively affects directional cell migration. Mechanistically, CEP85 and STIL can be recruited to the cell cortex in PLK4 kinase-dependent manner. Downregulation of CEP85 or STIL significantly reduces phosphorylation of ARP2 (also known as ACTR2), thus impairing actin reorganization and cell motility.

RESULTS AND DISCUSSION

CEP85 regulates directional cell migration

To identify additional components involved in PLK4-driven cell migration, we sought to examine the role of PLK4-associated centriole duplication factors in this process. To this end, we used siRNAs to deplete CEP192, CEP152, CEP85 and STIL in U-2 OS cells and measured the effect of their depletion on directional cell migration through conventional wound healing assays. Western blot analyses confirmed efficient depletion of each of these proteins in U-2 OS cells (Fig. S1A). Our results show that the downregulation of CEP85 in U-2 OS cells significantly suppressed wound healing to levels comparable to those observed upon PLK4 or CEP192 depletion (Fig. 1A,B). In agreement with these observations, we found that the reduction of PLK4, CEP192 or CEP85 levels exhibited a marked decrease in transwell cell migration (Fig. S1D–F). STIL depletion also prevented efficient wound closure and led to reduced transwell cell migration, although in a less-pronounced manner (Fig. 1A,B; Fig. S1D–F). Interestingly, although CEP152 is known to act as a scaffold for PLK4 activity (Sonnen et al., 2013; Park et al., 2014; Dzhindzhev et al., 2010), no noticeable defects in wound healing were observed upon its depletion, indicating that it does not act in this pathway. We next determined that the wound healing defects observed were not caused indirectly through impaired cell cycle progression, centrosome loss or positioning of the Golgi. Indeed, short-term treatment with centrinone B, a PLK4 inhibitor (Wong et al., 2015), significantly impaired wound healing (Fig. S1B,C), ruling out the possibility that centrosome loss was responsible for the observed defect. Consistent with this, depletion of PLK4, CEP192, CEP152, CEP85 or STIL under the low-serum conditions used in the migration assays, led to centrosome loss in ∼10% of cells (Fig. S2A,B), with no obvious defects in Golgi re-positioning in the direction of the wound (Fig. S2A,C) or cell cycle progression (Fig. S2D–K). These phenotypes are unlikely to be large enough to account for the cell migration defects observed upon CEP85 depletion.

Fig. 1.
Fig. 1.

CEP85 and STIL are required for directional cell migration. (A) Representative images from scratch-wound healing assays of U-2 OS cells transfected with different siRNAs. Scale bar: 400 μm. (B–D) Quantification (mean±s.d.; n=3 repeats for each of three independent experiments) of wound closure efficiency as shown in A as measured as the percentage of wound area closed (fraction of wound healed).

Previous work has established that CEP85 directly interacts with STIL to enable robust PLK4 kinase activation (Liu et al., 2018). To determine whether the CEP85–STIL interaction is also functionally important for PLK4-dependent cell migration (Rosario et al., 2015; Kazazian et al., 2017), we first depleted endogenous CEP85 or STIL in U-2 OS cells and examined the ability of siRNA-resistant wild-type (WT) CEP85 or STIL or the CEP85 Q640A+E644A or STIL L64A+R67A mutants (which compromise the CEP85–STIL interaction) to rescue cell migration (Liu et al., 2018). We observed that, compared to WT CEP85 or STIL, the expression of their mutants could not fully rescue the wound healing defect in CEP85- or STIL-depleted cells (Fig. 1C,D). These data suggest that a functional CEP85–STIL binding interface is required for efficient directional cell migration.

The binding of CEP85 to PLK4 but not CEP192 is required for cell migration

By undertaking BioID (proximity-dependent biotinylation), we and others have previously identified PLK4 and CEP192 as putative proximity interaction partners of CEP85 (Firat-Karalar et al., 2014; Liu et al., 2018). To build on these results, we conducted co-immunoprecipitation (co-IP) analysis and found that both WT CEP85 and the Q640A+E644A mutant could interact with PLK4, and that treatment with centrinone B had no effect on this association (Fig. 2A,B). These data suggest that the binding of CEP85 to PLK4 is independent of the CEP85–STIL interaction and PLK4 kinase activity. To determine which region in CEP85 is responsible for this interaction, we transiently expressed a set of CEP85 truncation mutants in U-2 OS cells for co-IP analysis. The results indicate that the middle region of CEP85 is responsible for its interaction with PLK4 (Fig. 2C,D). Intriguingly, the CEP85 ΔM mutant that was defective for PLK4 binding displayed reduced centriole localization, implying a potential role for this interaction in centriolar targeting of CEP85 (Fig. S3B). To further explore the putative CEP85–CEP192 interaction (Liu et al., 2018; Firat-Karalar et al., 2014), we performed a yeast two-hybrid assay and found that the N-terminal region of CEP85 can indeed associate with CEP192 (Fig. S3A). This putative interaction was not required for centrosome localization of CEP85 (Fig. S3B). Next, we set out to investigate the ability of the CEP85 ΔN and ΔM mutants to rescue centriole duplication and cell migration in U-2 OS cells with CEP85 depletion. Our results indicate that the expression of CEP85 ΔM mutant could not efficiently rescue these phenotypes compared to what was seen with the WT CEP85 and CEP85 ΔN mutant, indicative of a potential role for the CEP85–PLK4 interaction in cell migration and centriole assembly (Fig. 2E,F). To test this hypothesis, we sought to shorten their binding region and generated a set of truncation mutants in the middle region of CEP85 for co-IP analysis (Fig. S4A). We found that both the M1 and M2 regions can interact with PLK4 (although M2 appears to interact better with PLK4 than M1), implying that the entire M domain is important for robust PLK4 association (Fig. S4B). To functionally assess the effect of deleting the M2 region, we expressed an siRNA-resistant ΔM2 mutant in CEP85-depleted cells and found that it could not fully rescue the centriole duplication and cell motility phenotypes (Fig. S4C,D). Taken together, these results suggest that reducing the ability of PLK4 to interact with CEP85 negatively affects its ability to drive cell migration and centriole duplication. Furthermore, these findings could confirm the physiological relevance of the PLK4–CEP85 interaction.

Fig. 2.
Fig. 2.

CEP85 interacts functionally with PLK4. (A) Domain overview of human CEP85. cc, coiled-coil. The regions used in the vectors used in this work are indicated by the green lines. (B–D) Detection of expressed FLAG–CEP85 WT, Q640A+E644A mutant or fragments co-immunoprecipitating with Myc–PLK4. (E,F) U-2 OS cells expressing FLAG or the siRNA-resistant FLAG–CEP85 transgene were transfected with control or CEP85 siRNA for 72 h. (E) The graph indicates the percentage of cells with four centrioles at normal serum conditions (n=100 per experiment from three independent experiments). (F) The graph indicates the mean±s.d. percentage of the percentage of wound area closed (fraction of wound healed) at low serum conditions (n=3 repeats for each of three independent experiments). **P<0.01 (two-tailed t-test).

During directional cell migration, CEP192 has previously been shown to affect microtubule dynamics (O'Rourke et al., 2014). Consistent with this observation, western blot analysis indicated that CEP192 depletion led to a marked increase in the levels of the less-dynamic acetylated microtubules, while depletion of CEP85, STIL or PLK4 had no noticeable effect (Fig. S3C). These results suggest that the PLK4 interaction might be required for CEP85 to be able to drive centriole duplication and cell migration independently of CEP192.

CEP85 and STIL localize at the leading edge

We next sought to determine whether CEP85 and STIL can localize to the cell cortex. To test this, we transiently overexpressed mCherry–STIL in U-2 OS cells harboring tetracycline (Tet)-inducible GFP-tagged CEP85. Our results revealed that overexpressed WT CEP85 was able to colocalize with STIL at the cell cortex (Fig. 3A,B; Fig. S3D). In contrast, expression of the CEP85 ΔM and Q640A+E644A mutants failed to localize CEP85 and STIL at the cortex, suggesting that CEP85 binding to PLK4 and STIL is essential for their cortical localization (Fig. 3A,B; Fig. S3D).

Fig. 3.
Fig. 3.

CEP85 and STIL localization at the cell cortex. (A) Representative images of U-2 OS cells expressing GFP–CEP85 WT, ΔM, ΔN and Q640A+E644A mutants, and transfected with mCherry–STIL at low-serum conditions. (B) The graph shows the mean±s.d. percentage of cells with the described localization pattern (n=100 per experiment from three independent experiments). (C) U-2 OS cells expressing GFP–CEP85 WT and mCherry–STIL were either treated with centrinone B (1 μM) for 24 h or transfected with different siRNAs for 72 h at low serum conditions. (D) The graph shows the mean±s.d. percentage of cells with the described localization pattern (n=100 per experiment from three independent experiments). **P<0.01 (two-tailed t-test). Scale bars: 20 μm.

To test this hypothesis, we depleted endogenous STIL in U-2 OS cells using siRNA, and induced the expression of siRNA-resistant mCherry-tagged WT and mutant STIL to levels comparable to endogenous STIL (Fig. S3H). The results suggest that WT STIL still localizes to the cortex, whereas the L64A+R67A mutations perturb its cortical localization (Fig. S3F,G), suggesting that CEP85 binding is critical for the cortical localization of STIL. Next, we examined the effect of knocking down PLK4 and CEP192 on the cortical localization of CEP85 and STIL in U-2 OS cells. Our results indicate that the depletion of PLK4, or treatment with centrinone B, but not CEP192 depletion, perturbed the localization of CEP85 and STIL at the cortex (Fig. 3C,D; Fig. S3E). Together, our results suggest that PLK4 is an upstream regulator of CEP85 and STIL during cell migration.

CEP85 and STIL regulate the actin cytoskeleton

It has been shown that PLK4 phosphorylates ARP2 to mediate actin cytoskeletal rearrangement (Kazazian et al., 2017). Hence, we sought to determine whether CEP85 and STIL, which act to facilitate PLK4 activation (Liu et al., 2018), also contribute to this regulation. In this regard, we first used the canonical cell spreading assay in response to the stimulus of replating. Our results indicate that depletion of CEP85 and STIL adversely affected the cell spreading along with causing a significant cell size reduction, an effect that was comparable to PLK4 depletion or centrinone B treatment (Fig. 4A,B). We also confirm that the majority of those siRNA-transfected cells maintained their centrosomes in the spreading assay (Fig. S4F), suggesting that the phenotypes observed are independent of the centrosome. To further explore the molecular mechanism, we aimed to determine whether CEP85 and STIL impact PLK4-dependent phosphorylation of ARP2 (Kazazian et al., 2017). Using an antibody specific for phosphorylated ARP2, we observed that ARP2 phosphorylation at T237/T238 was significantly reduced in cells treated with centrinone B or depleted of PLK4, CEP85 and STIL (Fig. 4C,D). Taken together, these results indicate that the PLK4–CEP85–STIL module is required for ARP2/3 activation during directional cell migration (Fig. 4E).

Fig. 4.
Fig. 4.

CEP85 and STIL regulate ARP2/3-mediated actin reorganization. (A) Representative images of spreading assays in control, PLK4, CEP85 or STIL siRNA-transfected and centrinone B-treated U-2 OS cells. Cells were stained with Alexa Fluor 488–phalloidin. Scale bar: 20 μm. (B). The graph shows the quantification of relative cell area (mean±s.d.; n=100 per experiment from three independent experiments). (C) Western blot showing the levels of phospho- and total GFP–ARP2 in different conditions. (D) Quantification of fold change in phospho-ARP2 relative to total GFP–ARP2 (mean±s.d.; n=2 per experiment from six independent experiments). *P<0.05; **P<0.01 (two-tailed t-test). (E) A model for how PLK4–CEP85–STIL operates in the control of directional cell migration. CEP85 and STIL act downstream of PLK4, and their interaction facilitates PLK4 activation and subsequent ARP2 phosphorylation, which further regulates ARP2/3-mediated actin assembly and directed cell migration.

The work described here identifies an unexpected function of the centrosomal PLK4–CEP85–STIL module in directional cell migration. We find that PLK4 is required for the recruitment of CEP85 and STIL to the cell cortex, which is dependent on PLK4 kinase activity and its interaction with CEP85. This is consistent with our previous work indicating that the interaction between CEP85 and STIL is required for robust PLK4 kinase activation (Liu et al., 2018). This synergy promotes ARP2 phosphorylation in order to drive ARP2/3-dependent actin re-organization and directional cell migration. Overall, our data suggest that the PLK4–CEP85–STIL module plays a pivotal role in the regulation of both centriole duplication and cell motility.

It has emerged that centrosomes can act as actin-organizing centers through ARP2/3-based actin nucleation (Farina et al., 2016). The ARP2/3 complex consists of two actin related proteins (ARP2 and ARP3) and five accessory proteins, serving as unique nucleation sites for new actin filaments (Goley and Welch, 2006). Phosphorylation on residues T237, T238 and Y202 of ARP2 is required for stabilizing the ARP2/3 complex in an active conformation, which promotes efficient assembly of new actin (LeClaire et al., 2008). In the model proposed by Kazazian and colleagues, PLK4 is required for phosphorylation of ARP2 at the T237 and T238 residues, and ARP2 activity is critical for PLK4-driven cell migration (Kazazian et al., 2017). Our studies reveal the molecular basis of this regulation and characterize the CEP85–STIL complex as a novel modulator of PLK4-ARP2/3-mediated actin filament assembly and cell motility. Given the traditional view that these proteins functionally associate at the centrosomes and regulate centrosome-based actin nucleation, it must now also be considered that the PLK4–CEP85–STIL module can also control actin reorganization at the leading edge.

PLK4 is also implicated in the control of non-directional cell motility (Luo et al., 2019). In this model, CEP192 interacts with both PLK4 and AURKB. In response to exosome–WNT signaling stimulation, DVL2 initiates the recruitment of CEP192, PLK4 and AURKB to cell protrusions where PLK4 and AURKB act redundantly to drive formin-dependent actin reorganization (Luo et al., 2019). In contrast, as presented here, in the case of directional migration, PLK4 activity alone is required to drive the process. These observations raise the tantalizing possibility that other PLK4- and CEP192-associated components may be differentially regulated to control different types of cancer cell motility mediated by diverse actin nucleators, allowing for intricate contextual control of many forms of cell motility. Genetic amplification of PLK4 and its overexpression at the mRNA and protein level are frequently observed in such as breast, lung, pancreatic, rectum and stomach (Fig. S4E).

It will be important to dissect mechanistically, how the CEP85–STIL complex coordinates with PLK4 to activate ARP2/3-dependent actin polymerization and how this complex balances its centrosomal and non-centrosomal functions. Overall, our findings illuminate previously unappreciated regulatory mechanisms that participate in the control of directional cell migration.

MATERIALS AND METHODS

Cell culture

U-2 OS cells were grown under standard conditions, and were purchased from the ATCC. U-2 OS T-REx cells were cultured in McCoy 5A medium with 10% FBS, 2 mM GlutaMAX, zeocin (100 μg/ml) and blasticidin (3 μg/ml). U-2 OS T-REx cells with Tet-inducible Myc-tagged PLK4 were a kind gift Erich A. Nigg (Biozentrum, University of Basel, Switzerland), and were maintained in 10% FBS, 2 mM GlutaMAX and G418 (0.5 mg/ml). HCT116 cells were maintained in McCoy's 5A medium with L-glutamine (Life Technologies) supplemented with 10% FBS (Life Technologies) and 1% penicillin-streptomycin (Life Technologies). All human cell lines were cultured in a humidified 5% CO2 atmosphere at 37°C. All cell lines were tested and confirmed to be without mycoplasma contamination.

RNA interference

Luciferase duplex GL2 (5′-CGUACGCGGAAUACTTCG-3′) from Dharmacon was used as a control. The siRNAs against human PLK4 (M-005036-02-0005), CEP192 (L-032250-01-0005) and CEP152 (M-022241-01) were purchased from Dharmacon. CEP85 and STIL were depleted using the following siRNA oligonucleotide sequences: human CEP85 siRNA, 5′-CCUAGAGCAGGAAGUGGCUCAAGAA-3′ and human STIL siRNA, 5′-GCUCCAAACAGUUUCUGCUGGAAU-3′ (Liu et al., 2018). Transfections were performed using Lipofectamine RNAiMax (Invitrogen) according to the manufacturer's protocol.

Cloning and stable cell lines

GFP- or FLAG-tagged CEP85 fragments were recombined into the pcDNA5/FRT/TO vector backbone (Life Technologies) using a tetracycline-inducible CMV promoter. U-2 OS T-REx cells carrying full-length CEP85 and STIL or CEP85 Q640A+E644A and STIL L64A/R67A mutants were generated in our previous study (Liu et al., 2018). U-2 OS cells expressing the Tet-inducible FLAG-tagged siRNA-resistant CEP85 ΔM and ΔN transgenes were generated as previously described (Liu et al., 2018).

Wound healing assay

U-2 OS cells were seeded into 96-well plates and transfected with the indicated siRNAs and cultured at low serum concentrations (0.5% FBS). At 48 h post transfection, a single scratch wound was generated using the IncuCyte™ Wound Maker (Essen BioScience). Then cells were washed three times with PBS and cultured in serum-free medium for wounding healing assays. IncuCyte™ live-cell imaging systems (Essen BioScience) were utilized to conduct the 96-well scratch wound cell migration assay according to the manufacturer's protocol. Images were taken at 1 h intervals up to 24 h using a 10× objective lens. Image analysis was performed using ImageJ to measure the area of healed wound at t=6 h, 12 h, 18 h and 24 h.

For measuring the axis of nucleus–centrosome–Golgi, cells were fixed 0 or 6 h after generating wounds and stained with DAPI and antibodies specific to pericentrin (ab4448, dilution 1:500; Abcam) and GM130 (610823, dilution 1:400; BD Biosciences). The axis of nucleus–centrosome–Golgi was determined from the middle of the nucleus to the middle point between centrosome and Golgi. Angles between leading edge and axis of nucleus–centrosome–Golgi were measured with Image J. Radar graphs were drawn in RStudio (Ver 1.2.1335) with the ggplot2 library.

Transwell assay

U-2 OS cells (106) were seeded on six-well plates and then indicated siRNAs were transfected next day. At 1 day after siRNA transfection, cells were grown in McCoy's 5A medium containing 0.5% FBS for 24 h. For the migration assay, 5.0×104 cells were spread into the transwell (Millipore Sigma, CLS3422) with 0.1 ml of McCoy's 5A medium containing 0.5% FBS and then the transwells were inserted into 24-well plates containing 0.6 ml of McCoy's 5A medium supplemented with 10% FBS. After 24 h cells were fixed with 4% paraformaldehyde (PFA) for 2 min and cold methanol for 10 min. After removing cells inside the transwell, cells were stained with 0.5% Crystal Violet for 20 min. Cells were imaged on a Deltavision Elite DV imaging system (GE Healthcare) equipped with a sCMOS 2048×2048 pixels2 camera (GE Healthcare).

Spreading assays

U-2 OS cells were transfected with the indicated siRNAs and cultured at low-serum conditions. At 72 h post transfection, cells were trypsinized and re-plated onto 12-well plates with glass coverslips for another 6 h. Next, cells were fixed with 4% PFA and immunostained with Alexa Fluor 488–phalloidin (Invitrogen, A12379) for labeling F-actin. Images were acquired on a Deltavision Elite DV imaging system equipped with a 60×/1.4 NA objective and a sCMOS 2048×2048 pixels2 camera (GE Healthcare). Z-stacks (0.2 μm apart) were acquired, and images were deconvolved and projected using softWoRx (v6.0, Applied Precision). Quantification of cell area based on the F-actin signal was performed using ImageJ.

Immunofluorescence microscopy

Centrin and PCNT staining were performed following standard protocols with anti-centrin (clone 20H5, 04-1624, dilution 1:200; Millipore) and anti-pericentrin (ab4448, dilution 1:500; Abcam) antibodies as previously described (Liu et al., 2018). For the phalloidin staining, cells were fixed with 4% PFA at room temperature for 20 min and treated with permeabilization buffer (0.5% Triton X-100 in PBS) for another 10 min. Cells were then blocked in 1% BSA in PBS for 1 h and incubated with Alexa Fluor 488–phalloidin and DAPI in blocking solution for another hour. After a final washing step (three times with PBS for 5 min each), cells were inverted and mounted on glass slides with standard mounting solution (ProLong Gold antifade, Molecular Probes). Cells were imaged on a Deltavision Elite DV imaging system (GE Healthcare) equipped with a sCMOS 2048×2048 pixels2 camera (GE Healthcare). Z-stacks (0.2 μm apart) were collected, and images were deconvolved and projected using softWoRx (v6.0, Applied Precision).

Co-immunoprecipitation

The respective 293T or U-2 OS stable lines were seeded into 10-cm2 dishes and, 24 h later, transfected with 4 µg of plasmid DNA and incubated with tetracycline (2 μg/ml). At 48 h post transfection, transfected cells were washed with 1× PBS, then harvested and lysed immediately in lysis buffer [50 mM HEPES pH 8, 100 mM KCl, 2 mM EDTA, 10% glycerol, 0.1% NP-40, 1 mM DTT, and protease inhibitors (Roche)] for 30 min on ice. Lysates were frozen in dry ice for 5 min, then thawed and centrifuged for 20 min at 16,000 g at 4°C. Cleared supernatant was incubated with anti-FLAG M2 Affinity Gel (Sigma-Aldrich) for 3 h at 4°C. A fraction (10%) of the protein extracts were saved before the incubation with the beads (Inputs). After the incubation, the beads were pelleted and washed three times in lysis buffer. Samples (Inputs and IPs) were prepared by addition of Laemmli buffer and boiling at 95°C for 5 min. Immunopurified proteins were analyzed by immunoblotting with the indicated antibodies.

Western blotting

Cells were lysed in Laemmli sample buffer supplemented with a phosphatase inhibitor cocktail and benzonase nuclease. For phospho-ARP2 T237+T238 detection, cells were treated with 5.0 mmol/l pervanadate for 10 min before lysis. Proteins were loaded to 8% SDS-PAGE gels for electrophoresis and then electroblotted onto PVDF membranes (Immobilon-P, Millipore). Membranes were incubated with primary antibodies in Tris-buffered saline with 0.1% Tween-20 (TBST) in 5% skim milk powder or 5% BSA for the anti-ARP2 (against phosphor-T237+T238) antibody overnight at 4°C. Membranes were washed three times for 10 min each in TBST, and then incubated with secondary antibodies conjugated to horseradish peroxidase (HRP) for 1 h at room temperature. Western blots were developed using SuperSignal reagents (Thermo Fisher Scientific). Antibodies used in this study were: mouse polyclonal anti-CEP85 (H00064793-B01P, dilution 1:500; Abnova); mouse monoclonal antibodies anti-α-tubulin (clone DM1A, T6199, dilution 1:15,000; Sigma-Aldrich), anti-GFP (11814460001, dilution 1:2000; Roche), anti-FLAG (F3165, dilution 1:1000; Sigma), and anti-acetylated tubulin (T6793, dilution 1:2000; Sigma); rabbit polyclonal antibodies anti-CEP192 (A302-324A-1, dilution 1:1000; Bethyl Laboratories), anti-ARP2 phospho-T237+T238 (ab119766, dilution 1:500; Abcam) and anti-STIL (A302-441A, dilution 1:500; Bethyl Laboratories); and goat polyclonal antibody anti-c-Myc (ab19234, dilution 1:500; Abcam).

Flow cytometry

U-2 OS cells were cultured at low-serum conditions and harvested 72 h post-siRNA transfection from 10 cm plates. Fixation was performed using 4% PFA for 10 min at room temperature followed by permeabilization with 0.1% Triton-X for 15 min at room temperature. Cells were washed and resuspended in wash buffer (PBS plus 4% FBS). DAPI was added at a 1:100 final dilution, and cells were incubated in the dark at room temperature for 15 min. Flow cytometric analysis was performed on Fortessa X-20 (BD) using the UV laser for excitation and DAPI fluorescence was measured at 461 nm; 50,000 events were recorded for each sample. The doublet cell population was gated out and cell cycle distribution analysis was modeled using ModFit analysis, and the proportion of cells in G1, S and G2/M phases were quantified.

Statistical methods

Two-tailed unpaired Student's t-tests were performed for all P-values. Individual P-values, experiment sample numbers and the number of replicates for statistical testing are indicated in corresponding figure legends. Unless otherwise mentioned, all error bars are s.d., and the asterisk placeholders for P-values are *P<0.05 and **P<0.01.

Acknowledgements

We thank E. Nigg for the U-2 OS Tet-inducible PLK4 cells.

Footnotes

  • Competing interests

    The authors declare no competing or financial interests.

  • Author contributions

    Conceptualization: Y.L., L.P.; Methodology: Y.L., J.K., R.P.; Software: Y.L., V.S.; Validation: Y.L., J.K., V.S., M.C., M.v.B.; Formal analysis: Y.L., J.K., R.P.; Investigation: Y.L., M.v.B.; Resources: Y.L.; Data curation: Y.L.; Writing - original draft: Y.L., L.P.; Writing - review & editing: Y.L., J.K., R.P., V.S., M.C., J.M., M.v.B., L.P.; Supervision: L.P.; Project administration: L.P.; Funding acquisition: L.P.

  • Funding

    This work was supported by a Canadian Institutes of Health Research (CIHR) Foundation and Project Grants (#167279, MOP#130507 and 142192), ORF RE08-065, Krembil Foundation and CIHR Foundation grants to L.P. Y.L. was funded by a CIHR Doctoral Award.

  • Supplementary information

    Supplementary information available online at http://jcs.biologists.org/lookup/doi/10.1242/jcs.238352.supplemental

  • Received August 21, 2019.
  • Accepted February 21, 2020.

Peer review history

The peer review history is available online at https://jcs.biologists.org/lookup/doi/10.1242/jcs.238352.reviewer-comments.pdf

References

  1. ↵
    1. Ananthakrishnan, R. and
    2. Ehrlicher, A.
    (2007). The forces behind cell movement. Int. J. Biol. Sci. 3, 303-317. doi:10.7150/ijbs.3.303
    OpenUrlCrossRefPubMedWeb of Science
  2. ↵
    1. Arquint, C. and
    2. Nigg, E. A.
    (2016). The PLK4-STIL-SAS-6 module at the core of centriole duplication. Biochem. Soc. Trans. 44, 1253-1263. doi:10.1042/BST20160116
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Arquint, C.,
    2. Gabryjonczyk, A.-M. and
    3. Nigg, E. A.
    (2014). Centrosomes as signalling centres. Philos. Trans. R. Soc. Lond. B Biol. Sci. 369, 20130464. doi:10.1098/rstb.2013.0464
    OpenUrlCrossRefPubMed
  4. ↵
    1. Azimzadeh, J. and
    2. Marshall, W. F.
    (2010). Building the centriole. Curr. Biol. 20, R816-R825. doi:10.1016/j.cub.2010.08.010
    OpenUrlCrossRefPubMed
  5. ↵
    1. Basto, R.,
    2. Brunk, K.,
    3. Vinadogrova, T.,
    4. Peel, N.,
    5. Franz, A.,
    6. Khodjakov, A. and
    7. Raff, J. W.
    (2008). Centrosome amplification can initiate tumorigenesis in flies. Cell 133, 1032-1042. doi:10.1016/j.cell.2008.05.039
    OpenUrlCrossRefPubMedWeb of Science
  6. ↵
    1. Cunha-Ferreira, I.,
    2. Rodrigues-Martins, A.,
    3. Bento, I.,
    4. Riparbelli, M.,
    5. Zhang, W.,
    6. Laue, E.,
    7. Callaini, G.,
    8. Glover, D. M. and
    9. Bettencourt-Dias, M.
    (2009). The SCF/Slimb ubiquitin ligase limits centrosome amplification through degradation of SAK/PLK4. Curr. Biol. 19, 43-49. doi:10.1016/j.cub.2008.11.037
    OpenUrlCrossRefPubMed
  7. ↵
    1. De Pascalis, C. and
    2. Etienne-Manneville, S.
    (2017). Single and collective cell migration: the mechanics of adhesions. Mol. Biol. Cell 28, 1833-1846. doi:10.1091/mbc.e17-03-0134
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Dzhindzhev, N. S.,
    2. Yu, Q. D.,
    3. Weiskopf, K.,
    4. Tzolovsky, G.,
    5. Cunha-Ferreira, I.,
    6. Riparbelli, M.,
    7. Rodrigues-Martins, A.,
    8. Bettencourt-Dias, M.,
    9. Callaini, G. and
    10. Glover, D. M.
    (2010). Asterless is a scaffold for the onset of centriole assembly. Nature 467, 714. doi:10.1038/nature09445
    OpenUrlCrossRefPubMedWeb of Science
  9. ↵
    1. Farina, F.,
    2. Gaillard, J.,
    3. Guérin, C.,
    4. Couté, Y.,
    5. Sillibourne, J.,
    6. Blanchoin, L. and
    7. Théry, M.
    (2016). The centrosome is an actin-organizing centre. Nat. Cell Biol. 18, 65-75. doi:10.1038/ncb3285
    OpenUrlCrossRefPubMed
  10. ↵
    1. Firat-Karalar, E. N.,
    2. Rauniyar, N.,
    3. Yates, J. R., III. and
    4. Stearns, T.
    (2014). Proximity interactions among centrosome components identify regulators of centriole duplication. Curr. Biol. 24, 664-670. doi:10.1016/j.cub.2014.01.067
    OpenUrlCrossRefPubMed
  11. ↵
    1. Fung, E.,
    2. Richter, C.,
    3. Yang, H.-B.,
    4. Schäffer, I.,
    5. Fischer, R.,
    6. Kessler, B. M.,
    7. Bassermann, F. and
    8. D'angiolella, V.
    (2018). FBXL13 directs the proteolysis of CEP192 to regulate centrosome homeostasis and cell migration. EMBO Rep. 19, e44799. doi:10.15252/embr.201744799
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Godinho, S. A.,
    2. Picone, R.,
    3. Burute, M.,
    4. Dagher, R.,
    5. Su, Y.,
    6. Leung, C. T.,
    7. Polyak, K.,
    8. Brugge, J. S.,
    9. Théry, M. and
    10. Pellman, D.
    (2014). Oncogene-like induction of cellular invasion from centrosome amplification. Nature 510, 167-171. doi:10.1038/nature13277
    OpenUrlCrossRefPubMedWeb of Science
  13. ↵
    1. Goley, E. D. and
    2. Welch, M. D.
    (2006). The ARP2/3 complex: an actin nucleator comes of age. Nat. Rev. Mol. Cell Biol. 7, 713. doi:10.1038/nrm2026
    OpenUrlCrossRefPubMedWeb of Science
  14. ↵
    1. Gönczy, P.
    (2012). Towards a molecular architecture of centriole assembly. Nat. Rev. Mol. Cell Biol. 13, 425-435. doi:10.1038/nrm3373
    OpenUrlCrossRefPubMed
  15. ↵
    1. Guderian, G.,
    2. Westendorf, J.,
    3. Uldschmid, A. and
    4. Nigg, E. A.
    (2010). Plk4 trans-autophosphorylation regulates centriole number by controlling βTrCP-mediated degradation. J. Cell Sci. 123, 2163-2169. doi:10.1242/jcs.068502
    OpenUrlAbstract/FREE Full Text
  16. ↵
    1. Habedanck, R.,
    2. Stierhof, Y.-D.,
    3. Wilkinson, C. J. and
    4. Nigg, E. A.
    (2005). The Polo kinase Plk4 functions in centriole duplication. Nat. Cell Biol. 7, 1140-1146. doi:10.1038/ncb1320
    OpenUrlCrossRefPubMedWeb of Science
  17. ↵
    1. Holland, A. J.,
    2. Lan, W.,
    3. Niessen, S.,
    4. Hoover, H. and
    5. Cleveland, D. W.
    (2010). Polo-like kinase 4 kinase activity limits centrosome overduplication by autoregulating its own stability. J. Cell Biol. 188, 191-198. doi:10.1083/jcb.200911102
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Kazazian, K.,
    2. Go, C.,
    3. Wu, H.,
    4. Brashavitskaya, O.,
    5. Xu, R.,
    6. Dennis, J. W.,
    7. Gingras, A.-C. and
    8. Swallow, C. J.
    (2017). Plk4 promotes cancer invasion and metastasis through Arp2/3 complex regulation of the actin cytoskeleton. Cancer Res. 77, 434-447. doi:10.1158/0008-5472.CAN-16-2060
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Kleylein-Sohn, J.,
    2. Westendorf, J.,
    3. Le Clech, M.,
    4. Habedanck, R.,
    5. Stierhof, Y.-D. and
    6. Nigg, E. A.
    (2007). Plk4-induced centriole biogenesis in human cells. Dev. Cell 13, 190-202. doi:10.1016/j.devcel.2007.07.002
    OpenUrlCrossRefPubMedWeb of Science
  20. ↵
    1. Leclaire, L. L.,
    2. Baumgartner, M.,
    3. Iwasa, J. H.,
    4. Mullins, R. D. and
    5. Barber, D. L.
    (2008). Phosphorylation of the Arp2/3 complex is necessary to nucleate actin filaments. J. Cell Biol. 182, 647-654. doi:10.1083/jcb.200802145
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Levine, M. S.,
    2. Bakker, B.,
    3. Boeckx, B.,
    4. Moyett, J.,
    5. Lu, J.,
    6. Vitre, B.,
    7. Spierings, D. C.,
    8. Lansdorp, P. M.,
    9. Cleveland, D. W.,
    10. Lambrechts, D. et al.
    (2017). Centrosome amplification is sufficient to promote spontaneous tumorigenesis in mammals. Dev. Cell 40, 313-322.e5. doi:10.1016/j.devcel.2016.12.022
    OpenUrlCrossRefPubMed
  22. ↵
    1. Liu, Y.,
    2. Gupta, G. D.,
    3. Barnabas, D. D.,
    4. Agircan, F. G.,
    5. Mehmood, S.,
    6. Wu, D.,
    7. Coyaud, E.,
    8. Johnson, C. M.,
    9. Mclaughlin, S. H.,
    10. Andreeva, A. et al.
    (2018). Direct binding of CEP85 to STIL ensures robust PLK4 activation and efficient centriole assembly. Nat. Commun. 9, 1731. doi:10.1038/s41467-018-04122-x
    OpenUrlCrossRefPubMed
  23. ↵
    1. Luo, Y.,
    2. Barrios-Rodiles, M.,
    3. Gupta, G. D.,
    4. Zhang, Y. Y.,
    5. Ogunjimi, A. A.,
    6. Bashkurov, M.,
    7. Tkach, J. M.,
    8. Underhill, A. Q.,
    9. Zhang, L.,
    10. Bourmoum, M. et al.
    (2019). Atypical function of a centrosomal module in WNT signalling drives contextual cancer cell motility. Nat. Commun. 10, 2356. doi:10.1038/s41467-019-10241-w
    OpenUrlCrossRef
  24. ↵
    1. Mayor, R. and
    2. Etienne-Manneville, S.
    (2016). The front and rear of collective cell migration. Nat. Rev. Mol. Cell Biol. 17, 97. doi:10.1038/nrm.2015.14
    OpenUrlCrossRefPubMed
  25. ↵
    1. Moyer, T. C.,
    2. Clutario, K. M.,
    3. Lambrus, B. G.,
    4. Daggubati, V. and
    5. Holland, A. J.
    (2015). Binding of STIL to Plk4 activates kinase activity to promote centriole assembly. J. Cell Biol. 209, 863-878. doi:10.1083/jcb.201502088
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Nigg, E. A. and
    2. Raff, J. W.
    (2009). Centrioles, centrosomes, and cilia in health and disease. Cell 139, 663-678. doi:10.1016/j.cell.2009.10.036
    OpenUrlCrossRefPubMedWeb of Science
  27. ↵
    1. Ohta, M.,
    2. Ashikawa, T.,
    3. Nozaki, Y.,
    4. Kozuka-Hata, H.,
    5. Goto, H.,
    6. Inagaki, M.,
    7. Oyama, M. and
    8. Kitagawa, D.
    (2014). Direct interaction of Plk4 with STIL ensures formation of a single procentriole per parental centriole. Nat. Commun. 5, 5267. doi:10.1038/ncomms6267
    OpenUrlCrossRefPubMed
  28. ↵
    1. O'rourke, B. P.,
    2. Gomez-Ferreria, M. A.,
    3. Berk, R. H.,
    4. Hackl, A. M. U.,
    5. Nicholas, M. P.,
    6. O'rourke, S. C.,
    7. Pelletier, L. and
    8. Sharp, D. J.
    (2014). Cep192 controls the balance of centrosome and non-centrosomal microtubules during interphase. PLoS ONE 9, e101001. doi:10.1371/journal.pone.0101001
    OpenUrlCrossRef
  29. ↵
    1. Park, S.-Y.,
    2. Park, J.-E.,
    3. Kim, T.-S.,
    4. Kim, J. H.,
    5. Kwak, M.-J.,
    6. Ku, B.,
    7. Tian, L.,
    8. Murugan, R. N.,
    9. Ahn, M.,
    10. Komiya, S. et al.
    (2014). Molecular basis for unidirectional scaffold switching of human Plk4 in centriole biogenesis. Nat. Struct. Mol. Biol. 21, 696-703. doi:10.1038/nsmb.2846
    OpenUrlCrossRefPubMed
  30. ↵
    1. Paul, C. D.,
    2. Mistriotis, P. and
    3. Konstantopoulos, K.
    (2017). Cancer cell motility: lessons from migration in confined spaces. Nat. Rev. Cancer 17, 131. doi:10.1038/nrc.2016.123
    OpenUrlCrossRefPubMed
  31. ↵
    1. Rogers, G. C.,
    2. Rusan, N. M.,
    3. Roberts, D. M.,
    4. Peifer, M. and
    5. Rogers, S. L.
    (2009). The SCF(Slimb) ubiquitin ligase regulates Plk4/Sak levels to block centriole reduplication. J. Cell Biol. 184, 225-239. doi:10.1083/jcb.200808049
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Rørth, P.
    (2007). Collective guidance of collective cell migration. Trends Cell Biol. 17, 575-579. doi:10.1016/j.tcb.2007.09.007
    OpenUrlCrossRefPubMedWeb of Science
  33. ↵
    1. Rosario, C. O.,
    2. Kazazian, K.,
    3. Zih, F. S. W.,
    4. Brashavitskaya, O.,
    5. Haffani, Y.,
    6. Xu, R. S. Z.,
    7. George, A.,
    8. Dennis, J. W. and
    9. Swallow, C. J.
    (2015). A novel role for Plk4 in regulating cell spreading and motility. Oncogene 34, 3441. doi:10.1038/onc.2014.275
    OpenUrlCrossRefPubMed
  34. ↵
    1. Sonnen, K. F.,
    2. Gabryjonczyk, A.-M.,
    3. Anselm, E.,
    4. Stierhof, Y.-D. and
    5. Nigg, E. A.
    (2013). Human Cep192 and Cep152 cooperate in Plk4 recruitment and centriole duplication. J. Cell Sci. 126, 3223-3233. doi:10.1242/jcs.129502
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Strnad, P. and
    2. Gönczy, P.
    (2008). Mechanisms of procentriole formation. Trends Cell Biol. 18, 389-396. doi:10.1016/j.tcb.2008.06.004
    OpenUrlCrossRefPubMedWeb of Science
  36. ↵
    1. Wong, Y. L.,
    2. Anzola, J. V.,
    3. Davis, R. L.,
    4. Yoon, M.,
    5. Motamedi, A.,
    6. Kroll, A.,
    7. Seo, C. P.,
    8. Hsia, J. E.,
    9. Kim, S. K.,
    10. Mitchell, J. W. et al.
    (2015). Reversible centriole depletion with an inhibitor of Polo-like kinase 4. Science 348, 1155-1160. doi:10.1126/science.aaa5111
    OpenUrlAbstract/FREE Full Text
View Abstract
Back to top

Keywords

 Download PDF

Email
Short Report
Direct interaction between CEP85 and STIL mediates PLK4-driven directed cell migration
Yi Liu, Jaeyoun Kim, Reuben Philip, Vaishali Sridhar, Megha Chandrashekhar, Jason Moffat, Mark van Breugel, Laurence Pelletier
Journal of Cell Science 2020 133: jcs238352 doi: 10.1242/jcs.238352 Published 23 April 2020
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo
Citation Tools
Alerts

Article navigation

Other journals from The Company of Biologists

Advertisement

2020 at The Company of Biologists

Despite the challenges of 2020, we were able to bring a number of long-term projects and new ventures to fruition. While we look forward to a new year, join us as we reflect on the triumphs of the last 12 months.

Mole – The Corona Files

"This is not going to go away, 'like a miracle.' We have to do magic. And I know we can."

Mole continues to offer his wise words to researchers on how to manage during the COVID-19 pandemic.

Cell scientist to watch – Christine Faulkner

In an interview, Christine Faulkner talks about where her interest in plant science began, how she found the transition between Australia and the UK, and shares her thoughts on virtual conferences.

Read & Publish participation extends worldwide

“The clear advantages are rapid and efficient exposure and easy access to my article around the world. I believe it is great to have this publishing option in fast-growing fields in biomedical research.”

Dr Jaceques Behmoaras (Imperial College London) shares his experience of publishing Open Access as part of our growing Read & Publish initiative. We now have over 60 institutions in 12 countries taking part – find out more and view our full list of participating institutions.

JCS and COVID-19

For more information on measures Journal of Cell Science is taking to support the community during the COVID-19 pandemic, please see here.

If you have any questions or concerns, please do not hestiate to contact the Editorial Office.