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Radixin regulates cell migration and cell–cell adhesionthrough Rac1

Ferran Valderrama1,2, Subangi Thevapala2 and Anne J. Ridley1,*1Randall Division of Cell and Molecular Biophysics, King’s College London, New Hunt’s House, Guy’s Campus, London SE1 1UL, UK2Division of Biomedical Sciences, St George’s, University of London, Cranmer Terrace, London SW17 0RE, UK

*Author for correspondence (anne.ridley@kcl.ac.uk)

Accepted 8 March 2012Journal of Cell Science 125, 3310–3319� 2012. Published by The Company of Biologists Ltddoi: 10.1242/jcs.094383

SummaryThe ERM proteins ezrin, radixin and moesin are adaptor proteins that link plasma membrane receptors to the actin cytoskeleton. Ezrin andmoesin have been implicated in cell polarization and cell migration, but little is known about the involvement of radixin in these processes.

Here we show that radixin is required for migration of PC3 prostate cancer cells, and that radixin, but not ezrin or moesin, depletion byRNA interference increases cell spread area and cell–cell adhesion mediated by adherens junctions. Radixin depletion also alters actinorganization, and distribution of active phosphorylated ezrin and moesin. Similar effects were observed in MDA-MB-231 breast cancer

cells. The phenotype of radixin-depleted cells is similar to that induced by constitutively active Rac1, and Rac1 is required for the radixinknockdown phenotype. Radixin depletion also increases the activity of Rac1 but not Cdc42 or RhoA. Analysis of Rac guanine nucleotideexchange factors (GEFs) suggests that radixin affects the activity of Vav GEFs. Indeed, Vav GEF depletion reverses the phenotype of

radixin knockdown and reduces the effect of radixin knockdown on Rac1 activity. Our results indicate that radixin plays an important rolein promoting cell migration by regulating Rac1-mediated epithelial polarity and formation of adherens junctions through Vav GEFs.

Key words: ERM proteins, Rho GTPase, actin cytoskeleton, cadherin, cell migration, cell–cell adhesion

IntroductionThe ezrin/radixin/moesin (ERM) family of proteins act as

reversible linkers between cell surface receptors and the actincytoskeleton, and participate in signal transduction pathwaysregulating cell migration and cell polarity (Fehon et al., 2010).

They contribute to the formation of dynamic actin filament-containing membrane structures such as filopodia, uropods,lamellipodia and membrane ruffles, the cleavage furrow individing cells, and cell–cell adhesions (Bretscher et al., 2002;

Gautreau et al., 2002; Niggli and Rossy, 2008).

Ezrin, radixin and moesin are closely related ,80-kDa proteinsthat consist of three domains: an N-terminal FERM (four point

one-ezrin–radixin–moesin) domain linked to a C-terminal F-actin-binding domain through an a-helical domain (Fehon et al., 2010;Neisch and Fehon, 2011). The FERM domain interacts either

directly with membrane receptors such as CD44, CD43, andintercellular adhesion proteins (ICAMs) or indirectly throughscaffold proteins such as EBP50 (ezrin binding protein 50). It also

binds to phosphatidylinositol 4,5-bisphosphate [PtdIns(4,5)P2](Bretscher et al., 2002; Gautreau et al., 2000; Gautreau et al.,2002; Matsui et al., 1998; Nakamura et al., 1995; Oshiro et al.,1998). ERM proteins can exist in an inactive ‘closed’

conformation, where the C-terminal domain binds to the N-terminal FERM domain. In this conformation they are found in thecytosolic fraction of cells and the binding sites for plasma

membrane proteins and F-actin are masked. ERM proteins areactivated through conformational changes induced by binding toPtdIns(4,5)P2 and subsequent phosphorylation of a conserved C-

terminal threonine residue [T567 in ezrin, T564 in radixin andT558 in moesin] (Brown et al., 2003; Gautreau et al., 2000;Nakamura et al., 1995; Roch et al., 2010; Suda et al., 2011).

Several kinases, including members of the protein kinase C,

ROCK and MRCK families have been reported to regulate ERMfunction through phosphorylation of this threonine (Baeyens et al.,

2010; Belkina et al., 2009; Hebert et al., 2008; Matsui et al., 1998;Nakamura et al., 2000; Ng et al., 2001; Parisiadou et al., 2009;

Yamane et al., 2011).

ERM proteins can act both upstream and downstream of Rho

GTPases, which are key regulators of cell polarity and migration(Hall, 2009; Hughes and Fehon, 2007). Rho GTPases, cycle

between an active GTP-bound and an inactive GDP-boundconformation. They are activated by guanine nucleotide

exchange factors (GEFs) and inactivated by GTPase activatingproteins (GAPs). Some Rho GTPases are also regulated by binding

to GDP dissociation inhibitors (GDIs) or by phosphorylation(Hirao et al., 1996; Ivetic and Ridley, 2004; Mackay et al., 1997).

RhoGDI can associate with ERM proteins, thereby reducingRhoGDI interaction with GTPases and consequently promoting

their activation (Dovas and Couchman, 2005). Ezrin has been

reported to interact with PLEKHG6 (pleckstrin homology domaincontaining family G with RhoGEF domain member 6), a GEF for

RhoG and Rac1, to induce morphological changes in the apicalregion of epithelial cells (D’Angelo et al., 2007). In vitro studies

have also reported a direct interaction between the RhoGEF Dbland radixin (Prag et al., 2007; Takahashi et al., 1998). In this report

it is suggested that this interaction would compete with RhoGDI(Takahashi et al., 1998). On the other hand, RhoA and Rac

stimulate ERM activity by increasing the levels of PtdIns(4,5)P2

and/or by stimulating C-terminal phosphorylation, leading to

induction of microvilli enriched in activated ERM proteins(Auvinen et al., 2007; Shaw et al., 1997; Yonemura et al., 2002).

ERM proteins also bind to the cytoplasmic domain of the

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transmembrane receptor CD44 and this interaction is enhanced byRhoA activity (del Pozo et al., 1999; Lee et al., 2004; Tsuda et al.,2004). Finally, Rac1 activation in T-lymphocytes leads toinactivation of ERM proteins via dephosphorylation of the C-

terminal threonine (Cernuda-Morollon et al., 2010).

The expression of ERM proteins has been investigated in

several cancer types, including prostate cancer. For example,ezrin expression has been correlated with adverse prognosis,showing high levels of expression in both high grade prostate

intraepithelial neoplasia and prostate adenocarcinoma (Pang et al.,2004; Valdman et al., 2005). Androgen-induced cell invasionrequires hormonal regulation of ezrin phosphorylation in the

prostate cancer cell line, LNCaP (Chuan et al., 2006). Recently, astudy analysing radixin and moesin expression in normalprostate, high grade prostate intraepithelial neoplasia andprostatic adenocarcinoma tissues showed a slight decrease in

radixin levels in adenocarcinomas (Bartholow et al., 2011). Onthe other hand, radixin has been implicated in promoting breastcancer metastasis in an experimental metastasis model

(Valastyan et al., 2009).

Although the functions of ezrin and moesin have been

extensively studied in cell migration, a role for radixin has notbeen described. Here we show that radixin depletion severelyimpairs migration of prostate cancer PC3 cells. This inhibition is

accompanied by a change in cell shape and increased spread areaconcomitant with reorganization of the actin cytoskeleton, focalcontacts and cell–cell contacts. We demonstrate that Rac1, whichis activated by radixin depletion, mediates these changes. Our

results indicate that radixin levels regulate the transition from anon-migratory epithelial morphology with cell–cell adhesions toa migratory morphology by altering Rac1 activity.

ResultsRadixin regulates cell migration

To elucidate the role of radixin in cell migration, we depleted ERMprotein expression in prostate carcinoma-derived PC3 cells bytransient transfection with siRNA (sequences in supplementarymaterial Table S1). Radixin, ezrin and moesin were each

efficiently knocked down by 4 different siRNAs (Fig. 1A)Radixin depletion did not affect the levels of ezrin or moesin(Fig. 1B).

Cell migration was analysed by tracking cells in moviesgenerated by time-lapse microscopy (Fig. 1C; supplementary

material Movies 1 and 2). PC3 cells normally grow and migrateas single cells with a mesenchymal morphology, and migratewith a mean speed of 1.3460.04 mm/minute. In contrast, radixin-

depleted PC3 cells had a migration speed of approximately 38%of that of control siRNA-transfected cells, observed with 4independent siRNAs targeting radixin (Fig. 1D; supplementarymaterial Movies 1 and 2). The inhibition of migration following

radixin suppression appeared to be independent of the substrateas similar results were obtained on uncoated plastic and oncollagen-I or fibronectin-coated surfaces (data not shown).

Radixin-depleted cells were flatter with a more circular shapethan control cells; they also showed persistent membrane ruffling

around their whole periphery, rather than the small and localisedmembrane ruffles in discrete regions observed in control cells(Fig. 1E; supplementary material Movies 3 and 4). In addition,

we observed a fourfold increase in the cell spread area of radixin-depleted cells compared to control cells, either quantified fromfixed cells stained for F-actin (Fig. 1F) or from analysis of cells

in time-lapse movies (data not shown). In contrast, depletion ofmoesin or ezrin did not affect cell area (Fig. 1F). Although most

of the control cells had a migratory phenotype with a definedleading edge, we observed some cells in the whole populationwith a similar flat shape to radixin-depleted cells. We therefore

quantified the number of cells that had a circular, flat shape inboth control and radixin knockdown cells. We observed thatalmost 90% of the radixin knockdown cells were circular and flatbut just a 13% of control cells showed a similar phenotype

(Table 1).

Radixin regulates actin filament and focal contactdistribution

Since actin cytoskeleton and cell adhesion dynamics are required for

cell migration, we investigated whether radixin knockdown wasaccompanied by changes to the actin cytoskeleton and celladhesions. We observed that radixin-depleted PC3 cells had more

peripheral actin filament bundles than control cells, mainly localisedclose to the plasma membrane. This actin reorganization wasaccompanied by a re-localisation of the focal contact marker paxillin(Schaller, 2001; Schaller, 2004) from a uniform distribution around

the cell periphery to larger focal contacts that mainly localised at thetips of actin filament bundles (Fig. 2A). Active, C-terminallyphosphorylated ERM proteins (p-ERMs) often localise to

membrane ruffles and lamellipodia (Niggli and Rossy, 2008). Incontrol PC3 cells, p-ERMs were primarily localised to filopodia andother cell protrusions. In contrast, p-ERMs had a more punctate

distribution on the dorsal surface in radixin-depleted cells, similar tothe dorsal distribution of F-actin (Fig. 2B), a characteristic ofmicrovilli of epithelial cells (Fehon et al., 2010; Lan et al., 2006).This suggests that radixin-depleted PC3 cells have a more epithelial

phenotype. Quantification indicated that more than 80% of theradixin-depleted cells were positive for punctate F-actin and p-ERMin the top 20% of the cell height whereas only 12% of the control

cells had similar F-actin/p-ERM distribution at that height (Fig. 2C).Radixin therefore clearly regulates p-ERM distribution, although itdid not alter total levels of p-ERM, as determined by western

blotting (supplementary material Fig. S1).

Radixin depletion similarly induced cell spreading, increasedperipheral actin filament bundles and accumulation of p-ERM on

the dorsal surface in MDA-MB-231 breast cancer cells, indicatingthat the phenotype is not PC3-cell specific (supplementarymaterial Fig. S2).

Radixin depletion increases cell–cell contacts

From time-lapse movies, it was apparent that radixin-depletedcells formed more stable cell–cell contacts than control cells(Fig. 3A; supplementary material Movies 5 and 6). This wasquantified by determining the proportion of cells that maintained

contact with each other during the duration of each movie: over90% of radixin-depleted cells maintained contact with theirneighbour(s) over 16 h (Fig. 3B). In order to investigate in more

detail the basis for this radixin-regulated change in cell–celladhesion we analysed the distribution of adherens junctions,which are known to be important for establishing and

maintaining cell–cell contacts (Millan et al., 2010; Shapiro andWeis, 2009). PC3 cells express N-cadherin but not E-cadherin(supplementary material Fig. S3A), and radixin knockdown did

not induce E-cadherin expression (supplementary material Fig.S3B). In control PC3 cells N-cadherin was predominantlydistributed throughout the cytoplasm with some localisation at

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the plasma membrane. After radixin knockdown, N-cadherin

accumulated at cell–cell contact regions (Fig. 3C). Similarly, b-

catenin, which binds directly to cadherin cytoplasmic tails

(Shapiro and Weis, 2009; Sung et al., 2008), was localised to

sites of cell–cell contact in radixin-depleted cells, whereas in

control cells it was homogenously distributed throughout the

cytoplasm (Fig. 3C).

Although cell spreading in radixin-depleted cells would be

expected to increase cell–cell contact, very few control PC3 cells

that interacted with each other showed N-cadherin and b-catenin

Fig. 1. Radixin depletion inhibits migration and increases cell area. (A) PC3 cells were transfected with four different siRNAs targeting radixin, ezrin, moesin

or control siRNA, and lysed after 48 hours. Cell lysates were processed for further analysis by western blot using specific antibodies against radixin. Control: non-

transfected cells. Blots were re-probed with antibodies to MLC as a loading control. (B) Western blot analysis of ezrin, radixin and moesin expression in radixin-

depleted cells. PC3 cells were transfected with siRNA to radixin, lysed after 48, 72 and 96 hours and processed for western blot analysis using antibodies against

the indicated proteins. (C) Cells were transfected with radixin siRNA or control siRNA. After 48 hours cells were trypsinised, replated and analysed by phase-

contrast time-lapse microscopy. Images are shown from movies of control siRNA-transfected cells (top) or radixin siRNA-2-transfected cells (bottom). Similar

results were obtained with radixin siRNA-3 (not shown). The time after the start of the time-lapse movies (h) is indicated. Scale bar: 50 mm. (D) Quantification of

migration speed of PC3 cells transfected with four different siRNAs to radixin (1–4) or control siRNA. (E) High magnification and high contrast images of cells

from supplementary material Movies 1 and 2. Membrane ruffles are indicated by arrows in control cells and arrowheads in radixin knockdown cells. Scale bar:

10 mm. (F) Quantification of cell spread area, determined by staining for F-actin, of cells transfected with radixin siRNA-2, ezrin siRNA-2, moesin siRNA-2 or

control siRNA. The spread area is expressed relative to that of control cells. Scale bar: 10 mm. Values are means 6 s.e.m. from at least three independent

experiments. Analysis of variance (ANOVA) was performed to assess the significance of the results shown in each graph. Subsequently a Dunnet’s multiple

comparison test was used to determine whether the differences were significant between control and knocked down populations (***P,0.001).

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accumulation or colocalisation in the regions of contact.

Quantification showed that 79% of radixin-depleted cells had

cell–cell contacts that were positive for N-cadherin, whereas only

3.5% of the control cells showing a similar flat phenotype had N-

cadherin in the cell–cell contact regions (Table 1). The interaction

of control cells did not seem to depend on whether they had an

elongated or spread morphology, whereas for radixin knockdown

cells there was a significant correlation between the presence of N-

cadherin in cell–cell contacts and a spread phenotype. This

indicates that the spread phenotype observed in some control cells

is not likely to be related to that induced by radixin knockdown.

Interestingly, when cells were grown to very high confluence,

the radixin-depleted cells showed a significant increase in cell

height (Fig. 3D), indicating that cell–cell contacts are likely to be

Table 1. Quantification of cell–cell contacts in radixin-depleted cells

Elongated cells Flat cells

siRNA Proportion Cell–cell contacts Proportion Cell–cell contacts

Control 87% 5% 13% 3.5%Radixin 11% 1.8% 89% 79%

PC3 cells were transfected with siRNA-2 to radixin or control siRNA. After 72 hours, cells were fixed and stained for F-actin (to reveal their shape) and for N-cadherin (to reveal cell–cell contacts). Cells were quantified for their morphology: they were defined as elongated when they had a polarized morphology with afront and a back, or as flat when it was not possible to define a front and a back and the cell had a circular morphology; and for cell–cell contacts, defined aspositive staining for N-cadherin between two cells.

Fig. 2. Radixin affects p-ERM, actin cytoskeleton and cell adhesion distribution. (A) Redistribution of actin filaments and phosphorylated paxillin (p-

Paxillin) in radixin-depleted cells through transfection of siRNA-2 to radixin (similar results were obtained with radixin siRNA-3; not shown). PC3 cells were

fixed and stained for F-actin and with antibodies to phosphorylated paxillin. Phosphorylated paxillin localises at the tips of F-actin bundles in radixin-depleted

cells (magnification of boxed regions in far right panels). Scale bar: 20 mm. (B) Distribution of F-actin and phosphorylated ERM proteins (p-ERM) in radixin-

depleted cells through transfection of radixin siRNA-2 (similar results were obtained with radixin siRNA-3, not shown). Cells were stained for actin filaments and

with antibodies to p-ERM. Far right panels show magnifications of the boxed regions. Scale bar: 20 mm. (C) Quantification of F-actin and p-ERM staining in the

top of cells grown in high-density cultures. Cells were maintained at confluence for 48 hours. Images were obtained by confocal microscopy (z-stacks) and

quantification was performed analysing positive staining in those images for actin and p-ERM in the top 20% of the cell height. Values are means 6 s.e.m. from at

least three independent experiments. ANOVA was performed to assess the significance of the results. Subsequently, a Dunnet’s multiple comparison test was used

to determine whether the differences were significant between control and knockdown populations (***P,0.001).

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more mature. In these high-density cultures, the cadherin-

associated protein p120-catenin (Carnahan et al., 2010)

localised to cell–cell contacts of radixin-depleted cells but not

those of control cells (supplementary material Movies 7 and 8).

Effects of radixin depletion are mediated by Rac1

Rho GTPases are known to be important both for cell migration

and for formation of adherens junctions in epithelial cells (Parri

and Chiarugi, 2010; Popoff and Geny, 2009; van Duijn et al.,

2010). Since radixin knockdown inhibited migration and

enhanced adherens junction formation, we investigated whether

radixin affected Rho GTPase activity. Radixin depletion led to an

increase in Rac1 activity, but no change in Cdc42 or RhoA

activity was observed (Fig. 4A). Total levels of Rac1, Cdc42 and

RhoA were not changed. Consistent with a role for Rac1 in

mediating the phenotypic changes induced by radixin

suppression, expression of constitutively active Rac1 (GFP–

Rac1–L61) but not dominant negative Rac1 (GFP–Rac1–N17)

stimulated cell spreading and increased N-cadherin levels at sites

of cell–cell contact (Fig. 4B; supplementary material Fig. S4).

GFP–Rac1–L61 colocalised with N-cadherin at cell–cell contacts

(Fig. 4B).

Fig. 3. Radixin depletion increases adherens junctions. (A) PC3 cells cultured at low-density were transfected with siRNA-2 to radixin or control siRNA. The

images are from movies of control siRNA-transfected cells (top) and radixin siRNA-transfected cells (bottom). White arrows indicate an example of a cell group

that remains as such through the whole movie. Similar results were obtained with siRNA-3 to radixin (not shown). The time after the start of the time-lapse movies

(h) is indicated. Scale bars: 50 mm. (B) Quantification of cells keeping and/or establishing cell–cell contacts, determined from movies such as those shown in

supplementary material Movies 5 and 6, of cells that were establishing and keeping a colony-like structure (more than two cells together through the whole

movie). In both control and radixin knockdown cells the number of cells that remained as a colony through the whole movie (black portion of the column) and the

ones that scattered (grey portion of the column) was quantified and the percentage of each type was calculated. (C) PC3 cells were transfected with radixin siRNA-

2 or control siRNA and analysed after 72 hours. Cells were fixed and stained for N-cadherin and b-catenin. b-catenin and N-cadherin colocalise in cell–cell

contact regions in radixin-depleted cells (right panels and magnification of boxed region in far right panels). Scale bar: 100 mm. (D) Quantification of cell height

in high-density cultures. Cells were maintained at confluence for 48 hours and examined by confocal microscopy to determine the cell height. Five different fields

containing at least 30 cells from each of three independent experiments were quantified using the calibrated ruler of the 3-D tool from the ZEN software (Zeiss).

ANOVA was performed to assess the significance of the results. Subsequently, a Dunnet’s multiple comparison test was used to determine whether the differences

were significant between control and knockdown populations (***P,0.001).

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In order to determine whether Rac1 was required for the

phenotype of radixin-depleted cells, cells were sequentially

transfected with siRNAs to Rac1 then radixin, or radixin then

Rac1 (Fig. 4C). Rac1 suppression both prevented the increase in

cell spread area induced by radixin depletion and reverted the

radixin knockdown phenotype to that of control cells. The

Fig. 4. Morphological effects of radixin depletion are Rac1-dependent. (A) PC3 cells were transfected with siRNAs to radixin or control siRNA. Cells were

lysed after 72 hours, and Rac1, Cdc42 and RhoA activity were determined by pull-down assays. A representative western blot is shown of active and total Rac1,

Cdc42, and RhoA (left), and quantification of relative activity from three independent experiments (right graph). Values are means 6 s.e.m. Two-way

ANOVA was performed to assess the significance of the results. Subsequently, a Bonferroni’s multiple comparison test was used to determine whether the

differences were significant between control and knocked down populations for each one of the Rho GTPases (***P,0.001). (B) PC3 cells were transfected with

the constitutively active form (GFP–Rac1–L61) or the dominant-negative form (GFP–Rac1–N17) of Rac1, and 48 hours later were fixed and stained for actin

filaments and with antibodies to N-cadherin (magnifications of boxed regions are shown in far right panels). (C) PC3 cells were sequentially transfected with

siRNAs to Rac1 and radixin or vice versa. Cells were then fixed and stained for actin filaments. The left panel shows a representative image of control cells; the

middle panels show images of the first round of transfection with siRNA to either Rac1 (top) or radixin (bottom); the right panels show images of the cells after the

second round of transfection with siRNA to either radixin (top) or Rac1 (bottom). Scale bars: 100 mm.

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changes induced by radixin depletion are therefore dependent on

Rac1.

Radixin depletion acts through Vav GEF activation toregulate Rac1 and cell spreading

To determine how radixin affected Rac1 activity, we first tested

whether there was an interaction between radixin and regulators

of Rac1, including Dbl and RhoGDI, which have previously been

reported to interact with radixin in vitro (Takahashi et al., 1998),

and Tiam1, which is known to promote adherens junction

formation via Rac1 (Buongiorno et al., 2008). We could not

detect interaction between endogenous proteins in any case by

immunoprecipitation (data not shown). The GEF activity of Vav

GEFs is known to be activated by phosphorylation of a conserved

tyrosine (Y174) in their acidic regions, adjacent to the PH-DH

GEF domains (Lopez-Lago et al., 2000). Interestingly, we

observed an increase in Y174 tyrosine phosphorylation of Vav

GEFs only in radixin-depleted cells but not ezrin- or moesin-

depleted cells (Fig. 5A). In order to determine whether Vav GEFs

were required for the phenotype of radixin-depleted cells, cells

were transfected with siRNAs to radixin or Vav GEFs alone or

together (Fig. 5B). Vav depletion prevented the increase in cell-

spread area induced by radixin depletion, although Vav depletion

alone did not alter cell morphology.

The increase in Vav phosphorylation upon radixin depletion

suggests that Vav GEFs could be responsible for the increase in

Rac1 activity. Depletion of Vav GEFs significantly reduced the

increase in Rac1 activity induced by radixin depletion (Fig. 6).

No change in Rac1 activity was observed upon Vav GEF

depletion in the absence of radixin knockdown (Fig. 6).

Together, these results indicate that the Vav GEF activation

following radixin depletion is required for Rac1-mediated cell

spreading.

DiscussionCompared to ezrin and moesin, little is known of radixin

function. Here we show for the first time that radixin regulates

cell shape, migration and cell–cell adhesion, and that it acts by

altering Rac1 activity. The inhibition of migration in radixin-

depleted cells is accompanied by an increase in cell area and

cadherin-mediated contacts. Our results indicate that the different

effects of ezrin, moesin or radixin depletion are not simply

explained by tissue-selective expression of the 3 isoforms. Only

radixin depletion but not ezrin or moesin depletion has a

profound effect on cell morphology in PC3 and MDA-MB-231

cells, even though ezrin and moesin are both expressed.

Radixin depletion induces changes in PC3 cell shape and

adhesion similar to acquisition of a more epithelial morphology,

despite the fact that they do not re-express E-cadherin. However,

mesenchymal cells (including fibroblasts and astrocytes)

frequently form N-cadherin-mediated adherens junctions (Li

et al., 2001; Zhu et al., 2010), and thus the junctions formed in

radixin-depleted PC3 cells are likely to resemble these. The

changes induced by radixin depletion are dependent on Rac1 since

Rac1 knockdown inhibits the radixin knockdown-induced increase

of cell area, and expression of constitutively active Rac1 induces

Fig. 5. Radixin regulates Vav tyrosine phosphorylation. (A) PC3 cells were transfected with the indicated siRNAs to radixin, ezrin or moesin or control

siRNA. After 72 hours, cells were lysed and cell lysates were analysed by western blotting for radixin, ezrin, moesin, phospho-Tyr174 Vav and total Vav. Values

are means 6 s.e.m. from at least three independent experiments. ANOVA was performed to assess the significance of the results shown in each graph.

Subsequently a Dunnet’s multiple comparison test was used to determine whether the differences were significant between control and knockdown populations

(***P,0.001). (B) PC3 cells were transfected with siRNAs to radixin, Vav GEFs (Vav1, Vav2 and Vav3), both radixin and Vav GEFs, or control siRNA. After 72

hours, cells were fixed and stained for actin filaments. Representative images are shown. Scale bar: 10 mm.

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cell spreading and N-cadherin accumulation at cell–cell contacts.

The role of Rac1 in inducing and maintaining adherens junctions is

well established (Hage et al., 2009; Ray et al., 2007), and thus

increased Rac1 activity could explain the high level of adherens

junctions in radixin-depleted cells. Several RacGEFs have been

implicated in adherens junction assembly. For example, RacGEF

Tiam1 stimulates adherens junction assembly in MDCK epithelial

cells (Malliri et al., 2004). Vav GEFs have also been implicated in

adherens junction signalling to Rac1 (Fukuyama et al., 2006).

Both Tiam1 and Vav have been implicated downstream of

phosphoinositide 3-kinases in regulating adherens junctions

(Rivard, 2009). The increased Rac1 activity in radixin-depleted

PC3 cells might be due to Vav GEFs rather than Tiam1, since

tyrosine phosphorylation of Vav GEFs was increased following

radixin knockdown and they were required for the increase in cell

size and Rac1 activity observed upon radixin depletion.

Both ezrin and moesin are implicated in regulating cell–cell

adhesion, but radixin has not previously been linked to this

process. For example, in the mouse intestinal epithelium, ezrin

deficiency results in fusion of villi, accompanied by the

appearance of elongated cell–cell junctions (Saotome et al.,

2004). In Caenorhabditis elegans, the single ERM protein erm-1

is required for lumen formation and positioning of adherens

junctions (Gobel et al., 2004). In zebrafish, moesin and the

endothelium-specific VE-cadherin both regulate endothelial

lumen formation during development of blood vessels (Zhu

et al., 2010).

Radixin localises to bile canicular membranes in mouse liver,

and radixin-deficient mice accumulate high levels of bilirubin in

their blood, possibly because they are unable to secrete it into the

bile (Kikuchi et al., 2002). However, this phenotype is dependent

on the genetic background of the mice (Fukumoto et al., 2007),

and the mice grow normally and are fertile. The underlying

defect is therefore unclear, although it could relate to a decrease

in microvilli in the apical bile canalicular membranes (Kikuchi

et al., 2002). Radixin-null mice also become deaf due to

progressive degeneration of cochlear stereocilia, which are

actin filament-based apical projections on hair cells (Kitajiri

et al., 2004). Cadherin-23 mutations lead to deafness in humans

(Muller, 2008), and thus it would be interesting to know whether

radixin affects cadherin-23 localisation in the cochlea, given that

we observe that radixin regulates N-cadherin localisation in PC3

cells.

Increased ezrin expression is a well-known prognostic marker

for tumour aggressiveness in a variety of cancers, and influences

tumour survival and progression (Hunter, 2004). Moesin

expression is also upregulated in some cancers, correlating with

progression (Cui et al., 2009). So far little is known about radixin

in cancer, although interestingly it is a target of the microRNA

miR-31, which inhibits breast cancer metastasis in animal models

(Valastyan et al., 2009). Depletion of the miR-31 targets radixin,

RhoA and integrin a5 together phenocopies the effects of miR-31

expression in suppressing metastasis (Valastyan et al., 2010). The

relative contribution of each of these three genes is not known,

however. Our data are consistent with a role for radixin in

promoting cancer cell migration, which could contribute to

cancer progression.

Materials and MethodsCell lines and reagents

PC3 and DU145 prostate cancer cells were grown in RPMI containing 25 mM

Hepes and 2 mM glutamine supplemented with 10% FCS, 100 mg/ml

streptomycin, and 100 units/ml penicillin (5% CO2, 37 C). MDA-MB-231 breast

cancer cells were grown in DMEM containing 25 mM Hepes and 2 mM glutamine

supplemented with 10% FCS, 100 mg/ml streptomycin, and 100 units/ml penicillin

(5% CO2, 37 C). Antibodies were obtained from the following sources: radixin

from Sigma-Aldrich (cat. no. R3653); ezrin, moesin, phospho-ERM, MLC andphospho-paxillin from Cell Signalling (cat. no. 3145, 3142, 2466, 3672 and 2541,

respectively); N-cadherin, E-cadherin, p120-catenin, b-catenin from BD

Transduction Laboratories (cat. no. 610921, 610404, 610134 and 610154,

respectively); Rac1 clone 23A8 from Millipore (cat. no. 05-389); Cdc42 from

Millipore (cat. no. 07-1466); RhoA from Cytoskeleton (cat. no. ARH03-A); Vav

and phospho-Vav from Santa Cruz (cat. no. sc-132 and sc-16408-R, respectively);

GAPDH from Sigma-Aldrich (cat. no. G-8795); actin filaments were visualized

with Alexa-Fluor-546- or Alexa-Fluor-633-labelled phalloidin from Invitrogen

(cat. no. A22283 and A22284, respectively). Alexa Fluor 488 anti-rabbit, AlexaFluor 546 anti-mouse, and Alexa Fluor 633 anti-mouse secondary antibodies were

from Invitrogen (cat. no. A11008, A11103 and A21126, respectively). HRP-anti-

rabbit and HRP-anti-mouse from GE-Healthcare (cat. no. NA934 and NA931,

respectively). Except when specifically stated, all the chemical reagents were

obtained from Sigma-Aldrich.

siRNA and DNA transfections

All siRNAs were from Dharmacon (Thermo Scientific); esiRNA (a mixture of

siRNAs that all target the same mRNA sequence) for vav1, vav2 and vav3 were

from Sigma-Aldrich; their sequences are specified in supplementary material

Table S1. Cells were reverse transfected with 50 nM siRNA or esiRNA using

Lipofectamine 2000 (Invitrogen) in complete medium without antibiotics. Mediumwas changed to complete medium 14 hours after transfection and samples were

collected. For experiments involving radixin knockdown, radixin 2 siRNA (as

specified in supplementary material Table S1) was used unless otherwise

indicated.

Fig. 6. Vav GEFs mediate the radixin-dependent increase of Rac1

activity. PC3 cells were transfected as indicated with siRNAs to radixin,

siRNAs to Vav GEFs (Vav1, Vav2 and Vav3) or control siRNAs. Cells were

lysed after 72 hours, and Rac1 activity was determined by pull-down assays.

A representative western blot is shown of active and total Rac1 and

quantification of relative activity from three independent experiments (graph).

Values are means 6 s.e.m. ANOVA was performed to assess the significance

of the results. Subsequently, a Bonferroni’s multiple comparison test was used

to determine whether the differences were significant between each condition

(*P,0.05; ***P,0.001).

Radixin in cell migration and adhesion 3317

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For DNA transfection cells were transfected using indicated plasmids. 1 mg of

DNA in 100 ml of serum-free medium and 3 ml of Lipofectamine 2000

(Invitrogen) in 100 ml of serum-free medium were mixed within 5 minutes and

the mixture added into cells in medium without antibiotics for 8 hours before

replacement with normal growth medium.

Western blotting

Cell lysates for western blot analysis were prepared at the specified times after

treatment using lysis buffer (50 mM Tris-HCl pH 8, 0.5 mM EDTA, 150 mMNaCl, 1% Triton X-100 plus protease and phosphatase inhibitors cocktails).

Lysates were mixed with 26Final Sample Buffer (FSB; Invitrogen) and resolved

on 4–12% polyacrylamide gels (Invitrogen), transferred to nitrocellulose

membranes and blotted with the indicated antibodies.

Immunofluorescence

PC3 cells grown on 13-mm diameter coverslips were fixed with 4%

paraformaldehyde for 20 min. Cells were permeabilised with 0.1% Triton X-

100, blocked with blocking buffer (1% BSA, 2% FCS in PBS) and incubated with

the indicated primary antibodies diluted in blocking buffer for 1 hour. After three

washes with PBS, cells were incubated with fluorochrome-conjugated secondary

antibody for 45 minutes. After five washes with PBS, coverslips were mounted inslides using fluorescent mounting medium (Dako).

Time-lapse microscopy

Images of PC3 cells were acquired at 37 C and 5% CO2 using a fully motorized,multi-field time-lapse microscope (Eclipse TE 2000-E; Nikon) with a charge-

coupled device camera (ORCA; Hamamatsu Photonics). Images were acquired at

different intervals using a 206 or 406 objective. Images were acquired using

MetaMorph software (MDS Analytical Technologies), and cells were tracked

using ImageJ software (National Institute of Health). Time-lapse videos were

acquired for 16 to 24 hours with images taken at 10-min intervals. Analysis of cell

speed was performed using the Chemotaxis plug in from Integrated BioDiagnostics

GmbH (IBIDI).

Confocal microscopy

Images were acquired at room temperature using a confocal microscope (LSM 510

META; Carl Zeiss, Inc.) with three single photomultiplier tube detectors mounted

on a inverted microscope (AxioObserver Z1; Carl Zeiss, Inc.) using a 4061.3 NAoil immersion objective (Carl Zeiss, Inc.). Images were acquired using Zen

software (Carl Zeiss, Inc.) and processed using Zen and Photoshop (Adobe)

software. Image analysis for cell height and morphology was performed using Zen

and Image J software.

Rho GTPase activity assays

Cells treated with siRNA against radixin or with siRNA control were scrapped and

processed for the Rac/Cdc42 activation assay or the RhoA activation assay

according to the manufacturer’s instructions (Upstate/Millipore and Cytoskeleton

Inc, respectively) using the following lysis buffer: 50 mM Tris-HCl, pH 7.5; 1mM

EDTA; 500 mM NaCl, 10 mM MgCl2, 1% (v/v) Triton X-100; 0.5% sodium

deoxycholate; 0.1% SDS, 10% (v/v) glycerol; 0.5% (v/v) 2-mercaptoethanol;protease inhibitor cocktail complete (from Roche) and phosphatase inhibitor

cocktail II and IV (from Calbiochem). Total input and bound samples were

subjected to SDS-PAGE and immunoblot analysis to determine the total and GTP

bound Rac1, Cdc42 or RhoA levels.

AcknowledgementsWe are grateful to the members of the Ridley laboratory for theirinsights during the development of the manuscript and to theMicroscopy Facility at St George’s University of London for theirhelp during the acquisition of some of the images included in themanuscript.

FundingThis work was supported by Cancer Research UK [grant numberC6620/A8833 to A.J.R.]; the Ludwig Institute for Cancer Research(core funding to A.J.R.); the Bettencourt-Schueller Foundation (Prizefor the Life Sciences to A.J.R.); King’s College London British HeartFoundation Centre of Excellence [grant number RE/08/003]; and StGeorge’s University of London (start-up grant to F.V.).

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.094383/-/DC1

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