© 2015. Published by The Company of Biologists Ltd.

The prion protein inhibits monocytic cell migration by stimulating β1 integrin

adhesion and uropod formation

Dion D Richardson*1, Simon Tol*1, Eider Valle-Encinas1, Cayetano Pleguezuelos1,

Ruben Bierings2, Dirk Geerts3 and Mar Fernandez-Borja1

(1) Department of Molecular Cell Biology and (2) Department of Plasma Proteins,

Sanquin Research and Landsteiner Laboratory, Academic Medical Center, University

of Amsterdam, Amsterdam, The Netherlands; (3) Department of Pediatric

Oncology/Hematology, Erasmus University Medical Center, Rotterdam, The

Netherlands

* Equal contribution

Correspondence: Mar Fernandez-Borja, Department of Molecular Cell Biology,

Sanquin Research, Plesmanlaan 125, 1066CX Amsterdam, The Netherlands; phone:

+3120-5121220; e-mail: m.fernandez@sanquin.nl

Keywords: Prion protein, monocyte, migration, adhesion, RhoA, ERM

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JCS Advance Online Article. Posted on 9 July 2015

Abstract

The broad tissue distribution and evolutionary conservation of the GPI-anchored

protein PrP suggests that it plays a role in cellular homeostasis. Since integrin

adhesion determines cell behavior, the proposed role of PrP in cell adhesion may

underlie the various in vitro and in vivo effects associated to PrP loss-of-function,

including the immune phenotypes described in PrP-/- mice. We have investigated the

role of PrP in the adhesion and (transendothelial) migration of human (pro)monocytes.

We found that PrP regulates 1 integrin-mediated adhesion of monocytes.

Additionally, PrP controls cell morphology and migratory behavior of monocytes:

PrP-silenced cells show deficient uropod formation on immobilized VCAM and

display bleb-like protrusions on the endothelium. Our data further show that PrP

regulates ligand-induced integrin activation. Finally, we found that PrP controls the

activation of several proteins involved in cell adhesion and migration, including RhoA

and its effector cofilin as well as proteins of the ERM family. We propose that PrP

modulates 1 integrin adhesion and migration of monocytes through RhoA-induced

actin remodeling by cofilin and through the regulation of ERM-mediated membrane-

cytoskeleton linkage.

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Introduction

The cellular prion protein (PrP) is a glycosyl-phosphatidyl-inositol-anchored

glycoprotein that resides in detergent-resistant membrane domains. The conversion of

PrP to a misfolded insoluble isoform (PrP Scrapie) is at the onset of

neurodegenerative disorders known as transmissible spongiform encephalopathies

(Colby and Prusiner, 2011). However, its physiological function is largely unclear

(Biasini et al., 2012), having been involved in multiple cellular processes such as

survival, apoptosis, differentiation and growth (Linden et al., 2008). This variety of

functions together with the high evolutionary conservation and wide tissue expression

of PrP suggest that this protein has a role in cellular homeostasis.

PrP is highly expressed in the hematopoietic system, particularly in hematopoietic

stem cells and in mature mononuclear leukocytes (Isaacs et al., 2006). Accordingly,

PrP has been reported to have a role in several hematopoietic and immune cell

functions in vivo, such as the self-renewal of long-term repopulating hematopoietic-

stem cells (Kent et al., 2009; Zhang et al., 2006), macrophage phagocytosis (de

Almeida et al., 2005) and T-cell activation (Ballerini et al., 2006). However, the role

of PrP in the innate immune response has hardly been explored with the exception of

one study in which peritonitis was induced in PrP-/- mice with zymosan (de Almeida

et al., 2005). PrP-/- mice showed decreased numbers of neutrophils and larger numbers

of monocytes in peritoneal infiltrates. Although the basis for this phenotype was not

investigated, one possibility is that PrP has an intrinsic role in leukocyte migration out

of the circulation and/or into the peritoneum. In addition, several in vivo and in vitro

studies have shown that PrP participates in the regulation of cell-cell (Malaga-Trillo et

al., 2009; Mange et al., 2002; Morel et al., 2008) and cell-matrix adhesion (Loubet et

al., 2012; Schrock et al., 2009).

Leukocyte extravasation takes place through the binding of cytokine-activated

leukocytes to the endothelium, followed by their migration across the endothelium

and subsequently the vascular basement membrane (Nourshargh et al., 2010).

Leukocyte integrin binding to endothelial ICAM-1 and VCAM-1 receptors is crucial

for leukocyte arrest and firm adhesion to the endothelium, allowing leukocytes to

resist flow-induced detachment. This requires integrin clustering upon ligand binding

to increase avidity and strengthen adhesiveness through integrin association with the

actin cytoskeleton via adapter proteins (Alon et al., 2005). Migrating leukocytes

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polarize into a leading edge and a uropod. The uropod is an adhesive structure

containing cholesterol-rich membrane domains, 1 integrins and other adhesion

receptors (i.e. ICAM-1 and -3) as well as actin-membrane linker proteins of the ERM

(ezrin-radixin-moesin) family (Friedl and Weigelin, 2008). To migrate, leukocytes

need to detach and retract their uropod, a process regulated by the small GTPase

RhoA (Liu et al., 2002; Worthylake et al., 2001).

Here we have addressed the role of PrP in monocyte adhesion and migration and

show that PrP silencing impairs monocyte adhesion to 1 integrin ligands and to

endothelial cells while increasing their motility. We show that PrP regulates ligand-

induced integrin activation and provide evidence that ERM proteins and the RhoA-

cofilin pathway are involved.

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Results

PrP colocalizes with integrins at cholesterol-rich domains in monocytic cells

In vivo experiments in mice have shown that PrP regulates the recruitment of

monocytes to inflamed peritoneum and the phagocytosis of dead cells by

macrophages, suggesting a role for PrP in innate immunity (de Almeida et al., 2005).

However, the mechanism underlying these effects has not been explored. Here, we

first investigated the role of PrP in the adhesion and migration of the human pro-

monocytic cell line U937. This model has been used extensively to study monocyte

adhesion and CXCL12-induced migration (Chan et al., 2001; Hyduk and Cybulsky,

2009) and allows for gene expression manipulation by transfection and transduction

techniques.

U937 cells express PrP on their cell surface as assayed by flow cytometry (Figure 1A).

To determine whether PrP accumulates in cholesterol-rich membrane domains (‘lipid

rafts’), we performed a membrane flotation assay. PrP was primarily found in the

low-density fractions together with the lipid raft marker flotillin-1 (Figure 1B),

indicating that PrP resides in lipid rafts in U937 cells. Further analysis of PrP

distribution by confocal microscopy showed that a fraction of U937 cells displays

polarized PrP localization on the membrane (Figure 1C). This fraction increased from

~20% to ~50% upon CXCL12 stimulation. In polarized cells, both PrP and 1

integrins clustered together in membrane caps where they colocalized (Pearson’s

coefficient larger than 0.5) (Figure 1C). In contrast, PrP and 1 integrins failed to

colocalize when non-clustered and in non-polarized cells (Pearson’s coefficient

smaller than 0.5) (Figure 1C). PrP also colocalized with the lipid raft-resident protein

flotillin-1 in polarized cells (Figure S1). These data indicate that integrins are

clustered in a fraction of U937 cells in steady-state conditions and that, upon cell

polarization, PrP and 1 integrins are recruited to flotillin-1 lipid raft domains.

Both the localization of PrP in lipid rafts and its colocalization with integrins suggest

a role for PrP in the regulation of monocyte adhesion and migration. To investigate

this, we generated stable U937 cell lines with reduced PrP expression by transduction

with lentiviruses expressing three different shRNAs to human PrP. In parallel, stable

cell lines were generated by transduction with lentivirus expressing a non-targeting

shRNA (shCtrl-U937) as a negative control. Flow cytometry detection of surface PrP

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showed that shPrP-1 (complementary to the 3’UTR of PrP mRNA) was able to reduce

PrP expression to ~35%, while shPrP-2 and -3 (complementary to the PrP coding

sequence) reduced PrP expression to ~70% (Figure 1D). Considering the moderate

efficiency of shPrP-2 and -3 in silencing PrP expression, we continued our work using

stable shPrP-1-transduced cell lines (shPrP-U937 cells).

PrP silencing did not affect the proliferation rate of U937 cells as evidenced by

routine cell counting of shCtrl- and shPrP-U937 cell cultures (not shown).

Accordingly, staining of apoptotic cells with annexinV-FITC showed no significant

differences between shCtrl- and shPrP-U937 (0.3% and 0.2% positive cells,

respectively). In addition, there was no difference in CXCL12-induced activation of

the survival kinase Akt between control and PrP-silenced cells (not shown).

PrP regulates 1 integrin adhesion of monocytes

We used stable shCtrl- and shPrP-U937 cell lines to explore the effect of PrP

depletion on CXCL12-induced cell chemotaxis in a Transwell assay using FN-coated

filters. ShPrP-U937 cells showed a significant increase in chemotaxis when compared

with shCtrl-U937 cells (Figure 2A), suggesting that PrP is a negative regulator of

chemotaxis. To confirm this, we induced PrP-dependent signaling by cross-linking of

surface PrP with the SAF61 antibody, previously shown to trigger signal transduction

in T lymphocytes and in a neuronal cell line (Stuermer et al., 2004; Mouillet-Richard

et al., 2000). U937 cells incubated with SAF61 displayed impaired migration (Figure

2B), confirming the negative regulatory effect of PrP on monocytic cell migration and

supporting the specificity of the shRNA-mediated PrP silencing. These effects of PrP

on cell chemotaxis were not due to changes in CXCR4 surface expression or

CXCL12-induced receptor internalization as tested by flow cytometry (not shown).

We then investigated whether changes in U937 cell migration were paralleled by

changes in cell adhesion by recording electrical resistance at 32 kHz over time of cells

adhering to ECIS electrodes coated with integrin ligands (FN, Fc-ICAM-1 or Fc-

VCAM-1) or with bovine serum albumin (BSA) (Figure 2C-E). This assay is based on

the increase in electrical resistance when cells spread on the electrodes following cell

adhesion (Wegener et al., 2000). Addition of cells to the electrode arrays resulted in a

slight increase in electrical resistance, which was consistently higher for shCtrl-U937

cells compared to shPrP-U937 cells (Figure 2C), although differences were not

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significant (not shown). Comparatively, a large increase in resistance was observed

upon the stimulation of cell adhesion with PMA (Figure 2C). ShCtrl-U937 cells

seeded on FN-coated electrodes showed a larger increase in PMA-stimulated adhesion

than shPrP-U937 cells (Figure 2C). U937 cells express 1 (VLA 4 and 5) and 2

integrins (LFA-1 and Mac-1) (Prieto et al., 1994), and under our experimental

conditions, preferentially adhered to 1 integrin ligands (FN and VCAM-1) (Figure 2

D, E). Consistent with a role for PrP in the regulation of 1 integrin-mediated

adhesion, shPrP-U937 cells showed reduced adhesion to FN (Figure 2D) and Fc-

VCAM-1 (Figure 2E). Thus, PrP silencing impairs, 1 integrin-mediated adhesion of

monocytic cells. This reduced integrin-mediated adhesion may facilitate leukocyte

motility and migration, which could explain our findings.

To confirm that our findings also apply to primary monocytes, we transduced freshly

isolated human monocytes with shPrP-1. ECIS adhesion assays showed similar

differences in adhesion to Fc-VCAM between shCtrl- and shPrP-monocytes as for

U937 cells (Figures 2F and 2G). Similar results were obtained using the human T

lymphocyte cell line Jurkat lymphocytic cells (not shown).

To establish that the differences using the ECIS adhesion assay were due to

differences in adhesion strength and not in cell shape (i.e. spreading), we performed

conventional adhesion assays by seeding cells on Fc-VCAM-coated plates and

counting adherent cells after thorough washing (Figure S2).

Together, these data indicate that PrP regulates 1integrin-mediated adhesion of

monocytes and suggest that it may have a similar role in other leukocyte types.

PrP regulates monocyte adhesion to the endothelium under flow

To assess the effects of PrP silencing in monocyte-endothelium interactions under

near-physiological flow conditions we used parallel flow chamber assays. Mixed

suspensions of shCtrl- and shPrP-U937 cells (1:1), differently labeled with either

calcein Red-Orange AM or calcein Green were stimulated with either CXCL12 or

PMA. After stimulation, labeled cells were perfused over TNF-stimulated HUVEC,

while recording fluorescent and bright-field time-lapse movies (movie snapshot in

Figure 3A). Adherent cells were quantified in each video frame as described in

Materials and Methods. ShPrP-U937 cells adhered in lower numbers when compared

with shCtrl-U937 cells, both after cell stimulation with CXCL12 (Figure 3B) and with

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PMA (Figure 3C). Our data show a significant reduction in firm adhesion of shPrP-

U937 cells to the endothelium under flow in both conditions (Figure 3D: CXCL12;

Figure 3E: PMA). Next, we transduced freshly isolated human monocytes with shPrP-

1 or with shCtrl and quantified adherent monocytes to CXCL12-coated endothelium

in the same assay as described for U937 cells. Our data show that PrP silencing also

impairs firm adhesion of primary monocytes to the endothelium (Figure 3F). We

performed similar experiments with PMA-activated monocytes, however we obtained

no significant differences between the adhesion of shCtrl- and shPrP-monocytes. We

think this can be explained by the low numbers of adherent cells due to monocyte

activation and adhesion to the tubing of the flow system.

Finally, similar results were obtained using PrP-silenced Jurkat cells (Figure S3A).

These findings suggest that PrP positively regulates the firm adhesion of monocytes,

and possibly other leukocyte types, to inflamed endothelium under physiological

conditions of flow-induced shear stress.

PrP regulates cell shape and migratory behavior of U937 cells on immobilized

VCAM

Our data above show that PrP regulates monocyte chemotaxis (Figure 2). To further

analyze the effects of PrP silencing on cell migration, we examined the migratory

behavior of shCtrl- and shPrP-U937 cells on immobilized Fc-VCAM and CXCL12 by

time-lapse video recording. Migrating shCtrl-transduced U937 cells formed long and

persistent uropods and moved in a polarized fashion extending lamellipodia and

filopodia at the cell front (Figure 4A and supplementary Movie 1). In contrast, shPrP-

U937 cells failed to form long uropods (Figure 4A and supplementary Movie 1).

Manual tracking of migrating cells and subsequent calculation of cell velocity and

accumulated distance of cell trajectories showed that shPrP-U937 cells were faster

and covered longer distances than shCtrl-U937 cells (Figure 4A). Lack of significant

differences in directness suggests that PrP silencing does not affect cell polarization

(Figure 4A). These data corroborate our findings using Transwell chemotaxis assays

and suggest that PrP silencing enhances leukocyte motility through the

downregulation of integrin-mediated adhesion.

Both 1 integrins and phosphorylated ERM proteins localize to the uropod of

migrating leukocytes (Lee et al., 2004; Martinelli et al., 2013). Similarly, phospho-

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ERM proteins accumulated in the uropod partially colocalized with 1 integrins in

shCtrl-U937 cells migrating on immobilized VCAM-1 (Figure 4B). In contrast,

shPrP-U937 cells mostly lacked uropods and contained reduced levels of phospho-

ERM proteins, which localized in discrete spots and in thin membrane protrusions

(Figure 4B). Measurement of uropod length in cells migrating on Fc-VCAM-1

showed that the average length of the uropod was reduced from 26.5 µm to 14.5 µm

upon PrP silencing (Figure 4B). To test whether uropod loss could be rescued by

overexpression of PrP in shPrP-U937 cells, we took advantage of the fact that shPrP-1

targets the 3’-UTR of PrP mRNA allowing for rescue with human PrP cDNA. We

transfected cells with GFP-PrP or GFP cDNAs by electroporation and subsequently

selected GFP-positive cells by flow cytometry. Since GFP-PrP mainly localizes to the

cell middle plane (in the Golgi, not shown), and the signal was therefore low at the

bottom plane, we co-detected PrP with SAF32 in non-permeabilized cells to clearly

show distribution of PrP on the membrane. Uropod formation in PrP-silenced cells

was rescued by overexpression of GFP-PrP, but not GFP (Figure 4C). In addition, PrP

on the cell surface (detected with anti-PrP antibodies in non-permeabilized cells)

partially co-localized with endogenous 1 integrin (Pearson’s correlation coefficient =

0.65±0.01 (mean±SEM)), which corroborates our findings in suspension cells.

ERM proteins link membrane proteins to the actin cytoskeletal cortex (Fehon et al.,

2010) and active phosphorylated ERM proteins localize to and regulate formation of

the uropod in migrating lymphoblasts (Lee et al., 2004; Martinelli et al., 2013). We

observed that the intensity of phospho-ERM labeling appeared to be reduced in

shPrP-U937 cells (Figure 4B). To substantiate this observation, we assessed ERM

protein phosphorylation by immunoblotting in cells stimulated with CXCL12 or PMA

in a time-course experiment. As moesin is the predominant ERM isoform expressed

in leukocytes (Ivetic and Ridley, 2004), we detected this protein as control for equal

protein loading. CXCL12 induced ERM phosphorylation whereas PMA reduced

phospho-ERM levels in a time-dependent manner (Figure 5A). Consistent with the

role of ERM proteins in uropod formation, levels of phospho-ERM proteins were

significantly reduced in shPrP-U937 cells stimulated with CXCL12 and slightly

decreased after stimulation with PMA (Figure 5A). In addition, basal phospho-ERM

levels were 1.5-fold lower in PrP-silenced cells when compared to control. These data

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suggest that PrP regulates uropod formation in monocytic cells through the regulation

of ERM protein phosphorylation.

ERM proteins are known to link the uropod marker CD44 to the actin cytoskeleton

(Friedl and Weigelin, 2008; Fehon et al., 2010). To determine whether the reduction

in ERM protein activation observed in shPrP-U937 cells had any effects in CD44

distribution, we immunodetected CD44 and flotillin-1 in cells migrating on

immobilized Fc-VCAM. In shCtrl-U937 cells, CD44 was enriched in uropods

together with flotillin-1 (Figure 5B). In contrast, CD44 was more uniformly

distributed on the membrane of shPrP-U937 cells, albeit flotillin-1 still showed a

polarized distribution (Figure 5B). These data indicate that PrP silencing alters CD44

membrane distribution.

Next, we examined in detail the migratory behavior of shCtrl- and shPrP-U937 cells

arrested on the endothelium under near-physiological flow conditions as described

above. Leukocyte firm adhesion to the endothelium is enhanced by PMA treatment,

which stimulates integrin adhesion and consequently inhibits leukocyte motility

(Cinamon et al., 2001). As expected, PMA-stimulated shCtrl-U937 cells remained

mostly round and immotile after arrest and adhesion to the endothelium. In contrast,

PMA-stimulated shPrP-U937 cells crawled over the endothelium while forming

dynamic bleb-like membrane protrusions (Figure 6A, see Movies 2, 3 and 4 in

supplementary material). On average 30% of shPrP-U937 cells arrested on the

endothelium displayed dynamic protrusions in contrast to only 10% of shCtrl-U937

cells (Figure 6B). PMA-activated shPrP-monocytes also showed increased membrane

activity compared to shCtrl-monocytes (see movie 5 in supplementary material),

although differences were smaller (the percentages of adherent cells with membrane

protrusions were 38.5% for shCtrl-monocytes and 53.3% for shPrP-monocytes).

However, these data should be taken with caution due to the low number of PMA-

activated monocytes that were flown over the endothelium. Finally, this phenotype

appears not to be unique for monocytic cells since PrP-silenced Jurkat cells displayed

similar bleb-like motility on activated endothelium as U937 cells (Figure S3B).

Finally, we assessed the role of PrP in leukocyte transendothelial migration by

quantification of the number of shCtrl-and shPrP-U937 cells undergoing diapedesis

across TNF-activated CXCL12-coated endothelium under flow conditions

(Grabovsky et al., 2000). Predictably, a significantly higher percentage of shPrP-U937

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cells transmigrated across the endothelium when compared to control cells (Figure

6C). Thus, following arrest on the endothelium, PrP-silenced cells appear to have a

diminished capacity to establish firm adhesion, displaying increased motility together

with high membrane protrusive activity as well as increased diapedesis.

PrP regulates integrin activity

Integrin-mediated adhesion is regulated by conformational changes (affinity) as well

as by the number of integrin molecules engaged in ligand binding (avidity) (Carman

and Springer, 2003). Our data show that PrP silencing leads to reduced monocyte

adhesion to 1 integrin ligands. To gain insight into the molecular mechanism

underlying these effects, we first investigated whether PrP silencing impaired integrin

affinity upregulation in in cells stimulated or not with CXCL12, PMA or Mn2+ for 10

min. PMA and CXCL12 induce intracellular signaling that leads to integrin activation,

while Mn2+ binds to surface integrins triggering a signaling-independent

conformational change (Carman and Springer, 2003). Total and activated 1 on the

surface of shCtrl- and shPrP-U937 cells were assessed by flow cytometry. No

differences in surface expression of total or activated integrins were detected between

shCtrl- and shPrP-U937 cells (Figure 7A).To further examine the effect of PrP

silencing on stimulus-induced upregulation of high-affinity 1 integrins, we

stimulated cells with CXCL12, PMA or Mn2+ in the presence of soluble Fc-VCAM.

Cell-bound Fc-VCAM was subsequently detected using APC-labeled F(ab’)2 goat

anti-human IgG and flow cytometry analysis (Chan et al., 2003). shPrP-U937 cells

consistently bound less Fc-VCAM (Figure 7B). Direct integrin activation with Mn2+

induced higher Fc-VCAM binding and reduced the differences between shCtrl- and

shPrP-U937 cells (Figure 7B). These data suggest that PrP-silencing does not affect

the number of integrins on the cell surface but decreases ligand-induced integrin

activation. We assessed this further by using multimeric Fc-VCAM complexes. For

this, preformed soluble multimeric ligand complexes were generated by incubation of

Fc-VCAM with IgG F(ab’)2 goat anti-human IgG (Konstandin et al., 2006). This

assay was previously shown to detect both integrin affinity and avidity in leukocytes

based on the fact that polyclonal F(ab’)2 fragments recognize different epitopes of the

human Fc moiety of Fc-VCAM (Konstandin et al., 2006). The differences in ligand

binding between control and PrP-deficient cells were more pronounced when

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multimeric Fc-VCAM complexes were used (Figure 7C). The exacerbation of the

differences between control and PrP-silenced cells when using a multivalent ligand

suggests that PrP silencing primarily impairs ligand-induced integrin avidity.

PrP regulates RhoA activation and actin polymerization

Integrin avidity is enhanced upon increased lateral mobility of integrins on the plasma

membrane, which allows them to form microclusters (Bakker et al., 2012; Buensuceso

et al., 2003). Integrin binding to the actin cytoskeleton may impose constraints to the

lateral mobility of integrins and reduce avidity. In addition, PrP-silencing was

previously shown to stabilize actin filaments in neuronal cells through a RhoA-

dependent pathway (Loubet et al., 2012). Therefore, we explored RhoA activation in

shCtrl- and shPrP-U937 cells stimulated with CXCL12 or PMA. ShPrP-U937 cells

displayed significantly elevated GTP-RhoA levels at 2 and 5 min of CXCL12

stimulation (Figure 8A). In contrast to CXCL12, PMA stimulation decreased RhoA

activation. However, the levels of GTP-RhoA were also slightly elevated in PMA-

stimulated PrP-silenced cells when compared to control cells (Figure 8A).

RhoA regulates the actin cytoskeleton by inducing both actin polymerization and

actin microfilament crosslinking and remodeling. Therefore, we investigated actin

polymerization dynamics in CXCL12-stimulated suspension cells using a more

sensitive flow cytometry-based assay (Voermans et al., 2001). In agreement with

previous published data (Voermans et al., 2001), we observed a burst of F-actin after

0.5 min stimulation, followed by a decrease to unstimulated levels at 10 min (Figure

8B). PrP-silenced cells showed higher F-actin content at every time point (Figure 8B).

Low concentrations of cytochalasin B are reported to enhance actin remodeling, while

at high concentrations, cytochalasin B causes actin cytoskeleton depolymerization. To

test whether altered actin dynamics underlies the adhesion defects of PrP-silenced

cells, we performed an ECIS adhesion assay in the presence of tow concentrations of

cytochalasin B. At 1 M, cytochalasin B increased adhesion of shPrP-U937 cells to

control levels (Figure 8C). At 5 M, cytochalasin B, reduced PMA-induced adhesion

of both shCtrl and shPrP-U937 cells (Figure 8C). These data suggest that reduced

actin dynamics mediates the effects of PrP silencing in cell adhesion.

Previously, the RhoA-regulated actin-severing protein cofilin was involved in the

regulation of actin microfilament stability by PrP in neurons (Loubet et al., 2012). To

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determine whether this pathway is operational in monocytes, we determined the levels

of phosphorylated (inactive) cofilin in cells adherent to immobilized Fc-VCAM. We

found that PrP-silencing increases phospho-cofilin levels (Figure 8D) and thus

inactivates cofilin. Together, these findings provide evidence suggesting that PrP

regulates monocyte adhesion and migration through the regulation of actin

remodeling by the RhoA-cofilin pathway.

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Discussion

Leukocyte migration across the vascular wall is of crucial importance for the immune

response. PrP has been previously involved in the influx of immune cells into the

peritoneal cavity during zymosan-induced peritonitis, where monocytes were

recruited in larger numbers in PrP-/- mice compared to wild type controls (de Almeida

et al., 2005). Our study now provides evidence for the molecular mechanisms

underlying the effects of PrP silencing on monocyte adhesion and migration, and

corroborate the immunomodulatory role of PrP.

Our findings show that PrP silencing increases chemotaxis and motility of monocytic

U937 cells on 1 integrin ligands in static conditions as well as on endothelium in

near-physiological flow conditions. In addition, we show that PrP-deficient U937

cells are able to transmigrate more efficiently than control cells. In contrast, activation

of endogenous PrP signaling by antibody-mediated crosslinking of surface PrP

impairs U937 cell chemotaxis. Collectively, these data suggest that PrP is a negative

regulator of monocyte (transendothelial) migration.

Leukocyte migration is dictated by the strength of integrin-mediated cell adhesion.

Although leukocyte attachment to the endothelial surface is required to resist flow-

induced shear stress forces, excessive adhesion abrogates leukocyte motility on the

endothelium as well as diapedesis across the endothelial monolayer (Cinamon et al.,

2001). We show that PrP silencing impairs pro-monocyte adhesion to 1 integrin

ligands (FN and VCAM-1) in static conditions, as well as to TNF-activated

endothelial cells under flow. In addition, we show that primary monocytes and Jurkat

cells also display defective adhesion and enhanced motility upon PrP silencing,

suggesting that PrP regulates 1 integrin-mediated interactions of monocytes (and

probably T cells) with the endothelium in vivo.

The regulation of integrin-mediated adhesion takes place through changes in affinity,

avidity and/or trafficking (Margadant et al., 2011). We observed no changes in

integrin affinity in PrP-silenced cells by using either an antibody specific for the high

affinity conformation of 1 integrins or a soluble 1 integrin ligand (Fc-VCAM1).

Likewise, we detected no differences between control and PrP-silenced cells in 1

integrin internalization by using an antibody-feeding assay (results not shown).

However, PrP silencing impaired the binding of multimeric Fc-VCAM complexes to

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cells. These data suggest that PrP controls 1 integrin adhesiveness through the

regulation of ligand-induced changes in integrin activation, likely through the

regulation of integrin avidity, which is determined by the number of integrins engaged

in ligand binding (Carman and Springer, 2003). VLA-4 (41) integrin

microclustering can induce adhesiveness by increasing valency, independent of

changes in integrin affinity (Grabovsky et al., 2000). The attachment of the integrin

cytosolic tail to the actin cytoskeleton restraints the lateral mobility of integrins on the

membrane and consequently impairs microclustering (Bakker et al., 2012;

Buensuceso et al., 2003). Thus, local microfilament remodeling might be required to

allow disengagement of integrins from cortical actin and allow them to cluster,

therefore increasing the strength of ligand binding (Bakker et al., 2012; Worthylake

and Burridge, 2003). We show here, that PrP silencing increases CXCL12-induced

RhoA activation as well as F-actin content. Interestingly, while CXCL12-induced

actin polymerization follows similar kinetics in shCtrl- and shPrP-U937 cells, actin

depolymerization is slower in PrP-silenced cells (Figure 8). Notably, increasing actin

depolymerization rates with low concentrations of cytochalasin B abrogate the effects

of PrP silencing on cell adhesion. Consistent with this, we found that PrP silencing

induces the inactivation of the actin-severing protein cofilin. Inactivation of cofilin by

phosphorylation on Ser3 is mediated by LIM-kinase which in turn is activated by

RhoA (Bravo-Cordero et al., 2013). In view of the above findings, we propose that the

stabilization of actin filaments by RhoA-induced cofilin inactivation prevents the

release of activated integrins from the cytoskeleton in PrP-deficient cells, therefore

impairing integrin microclustering and adhesiveness. Our findings are in agreement

with previously reported data showing increased cytoskeletal stability, RhoA

activation and cofilin inactivation in PrP-deficient neuronal cells (Loubet et al., 2012).

Interestingly, overexpression of PrP in primary rat hippocampal neurons induces

cofilin activation and the formation of actin-cofilin rods (Walsh et al., 2014). These

structures are found in Alzheimer disease brain and are shown to impair synaptic

function (Minamide et al., 2000). It is tempting to speculate that cofilin regulation is

critical both for the physiological function of PrP and for prion-induced

neurodegeneration.

We found that PrP colocalizes with integrins in pre-formed flotillin-1 domains

(Langhorst et al., 2006). In contrast to T cells, where PrP recruitment to flotilin-1

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membrane caps was induced by antibody-mediated PrP crosslinking (Stuermer et al.,

2004), we observed that PrP co-clustered with integrins in a fraction of U937 cells in

the absence of stimulation or antibody-mediated crosslinking. This suggests that a

percentage of U937 cells are ‘primed’ in resting conditions, which could explain the

differences observed in F-actin content and ligand binding in unstimulated cells.

Despite the colocalization of PrP and 1 integrin in flotillin-1 domains, we were

unable to show association of PrP and 1 in co-immunoprecipitation assays (data not

shown), albeit 1 integrin was previously identified as a PrP-interacting partner in a

proteomic study (Watts et al., 2009). We found that PrP silencing dramatically altered

the morphology and migratory behavior of cells moving on 2D surfaces. ShCtrl-U937

cells migrating on immobilized VCAM accumulated 1 integrins, phospho-ERM,

flotillin-1 and CD44 in the uropod, all bona fide uropod markers (Shulman et al.,

2009; Gomez-Mouton and Manes, 2007). In contrast, PrP-depleted cells generally

failed to form uropods and displayed a round shape, although 1 integrin and flotillin-

1 still showed a polarized distribution. Flotillins are essential for uropod formation by

neutrophils (Ludwig et al., 2010). Although PrP colocalizes with flotillin in U937

cells, flotillin distribution is not affected upon PrP silencing, which suggests that PrP

does not regulate uropod formation through the disruption of flotillin domains.

The uropod is an adhesive structure that need to be retracted during migration (Liu et

al., 2002; Smith et al., 2003; Worthylake et al., 2001). In agreement with this, PrP-

deficient cells migrated faster than control cells. We found that PrP silencing alters

the activation of two proteins involved in uropod dynamics: ERM and RhoA. First,

PrP silencing reduces ERM protein phosphorylation. ERM proteins are activated by

phosphorylation of a conserved C-terminal threonine residue, which enables them to

crosslink membrane receptors with the underlying actin cytoskeleton (Fehon et al.,

2010). ERM activation is crucial for uropod formation (Ivetic and Ridley, 2004; Lee

et al., 2004; Martinelli et al., 2013) and for 1 integrin-dependent T cell adhesion and

polarization (Chen et al., 2013; Liu et al., 2002). Therefore, the inhibition of ERM

phosphorylation in the absence of PrP likely contributes to both the lack of uropods

and the reduced 1 integrin adhesion observed for PrP-deficient monocytes.

RhoA is crucial for uropod de-adhesion and retraction during migration (Liu et al.,

2002; Smith et al., 2003; Worthylake et al., 2001). In neutrophils, activation of

myosin by RhoA has been implicated in actomyosin-driven uropod retraction (Eddy

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et al., 2000). However, we found that phospho-myosin levels were equal in shCtrl-

and shPrP-U937 and they did not increase by stimulating migration (data not shown).

This suggests that actomyosin contraction does not play a role in uropod retraction in

monocytic cells. In line with this, myosin inhibition did not affect uropod retraction in

migrating monocytes (Worthylake et al., 2001).

PMA-dependent integrin activation induces leukocyte firm adhesion to the

endothelium and blocks motility and migration (Cinamon et al., 2001). Thus

‘excessive’ integrin activation prevents migration. In line with this, PMA-stimulated

control cells firmly adhered to the endothelium, while PMA-stimulated PrP-deficient

cells crawled over the endothelium while protruding dynamic bbleb-like extensions.

This motility could be prompted by the weaker adherence of PMA-simulated PrP-

silenced cells. In addition, decreased membrane tension due to lower levels of active

ERM proteins may also contribute to this phenotype. In support of this, constitutively

active ezrin was shown to increase lymphocyte membrane tension, reduce chemokine-

induced shape changes and block transendothelial migration (Liu et al., 2002).

In summary, we propose that PrP has a role in integrin-mediated adhesion and

motility through the regulation of ERM membrane-cortex adaptors and the RhoA-

cofilin pathway that controls cytoskeleton remodeling. This may be a general

mechanism for the reported regulation of cell-cell and cell-matrix adhesion by PrP in

different cell types (Petit et al., 2013). Also, our data contributes to understand

additional roles of PrP in neurodegeneration.

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Methods

Cell culture and lentiviral transduction

Human pro-monocytic U937 cells were transduced with lentivirus containing the

validated Sigma MISSION shRNA plasmids targeting human PrP (PrP-1,

TRCN0000083488; PrP-2, TRCN0000083490; PrP-3, TRCN0000083491) or non-

targeting shRNA (Ctrl, SHC002) as a negative control. Subsequently shRNA-

expressing cells were selected by culturing in RPMI medium containing 10% fetal

calf serum and 1.5 µg/mL puromycin. Before each experiment, cells were cultured in

puromycin-free medium for at least 24 h. PrP expression in control and knock-down

cell lines was tested before each experiment by flow cytometry with SAF32 anti-PrP

antibody. HUVEC (Primary Human Umbilical Vein Endothelial Cells) were cultured

in Endothelial Growth Medium-2, both from Lonza.

Human monocytes were isolated from PBMCs by positive selection with magnetic

CD14-beads (Milteny Biotech). For transduction, shRNA lentiviruses were incubated

with 4 g/mL Polybrene for 10 min at RT prior to their addition to monocyte

suspensions (2x106 cells/mL) in complete RPMI. M-CSF was added at a

concentration of 10 ng/mL to keep monocytes steady without inducing differentiation

(Troegeler et al., 2014). After 48 h, PrP expression was analyzed by flow cytometry.

The maximum efficiency of PrP knock-down was of ~30-60%. Only those monocytes

with at least 50% knock-down were used for experimental assays.

Antibodies and reagents

Mouse monoclonal antibodies (Abs) against PrP (SAF32 and SAF61) were from Spi-

Bio. Anti-1 integrin Ab (clone P5D4) was from Abcam. Activated 1 integrins were

detected with HUTS-4 (Millipore) and Mab24 (Abcam), respectively. Flotillin-1

rabbit polyclonal Ab was from Abcam and transferrin receptor mouse monoclonal Ab

from Life Technologies. RhoA monoclonal mouse Ab was from Santa Cruz

Biotechnology. Rabbit antibodies against phospho-ERM proteins (ezrin (Thr567),

radixin (Thr564), moesin (Thr558)), phospho-cofilin (Ser3), cofilin and myosin light

chain (MLC) as well as mouse monoclonal anti phospho(Ser19)-myosin light chain

were from Cell Signaling. Mouse monoclonal anti-moesin was from BD Biosciences.

FITC-labeled anti-CXCR4 mouse monoclonal was from R&D Systems. Mouse

monoclonal anti-actin was from Sigma. HRP- and Alexa dye-conjugated secondary

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Abs were from Dako and Life Technologies, respectively. Human fibronectin was

from Biopur. Recombinant human Fc-ICAM and Fc-VCAM were from R&D

Systems. (APC-labelled) polyclonal goat anti human IgG F(ab’)2 were from Jackson

ImmunoResearch Laboratories. AnnexinV-FITC was from Pharmingen. Human

recombinant TNF and CXCL12 were from Peprotech. Phorbol-12-myristate-13-

acetate (PMA) and cytochalasin B were from Sigma. Calcein Green AM and calcein

Red-Orange AM were from Life Technologies.

Membrane flotation assay

20x106 cells were lysed on ice for 15 min in TXNE buffer containing 50 mM Tris-

HCl (pH 7,4); 150 mM NaCl; 5 mM EDTA; 1 mM DTT; 1% Triton X-100 and

cOmplete Protease Inhibitor Cocktail Tablets (Roche). Lysates were mixed with 1.8

mL of 60% OptiPrep (Sigma) in TXNE buffer. Subsequently, 7.5 mL of 28%

OptiPrep and 1.8 mL of TXNE buffer were layered on top. Gradients were

centrifuged at 200,000 g for 3 h at 4°C. 1-mL fractions were taken from the top of the

gradient and proteins were precipitated for 30 min on ice by addition of an equal

volume of 20% trichloroacetic acid and pelleted at 10,000 g for 15 min at 4°C. Pellets

were washed with cold acetone and solubilized in SDS sample buffer.

Chemotaxis

Transwell chemotaxis assays were performed and analyzed as previously described

(Lorenowicz et al., 2006). Crosslinking of surface PrP was performed by incubating

cells for 30 min at 37°C with 0.5 µg/mL of anti-PrP SAF61 Ab.

ECIS adhesion assays

Cell adhesion to fibronectin (FN), BSA, Fc-VCAM or Fc-ICAM was measured using

Electrical Cell-Substrate Impedance Sensing system (ECIS, Applied Biophysics).

ECIS 8W10E electrode arrays were coated with 20 µg/mL FN or BSA during 16 h at

4°C. For Fc-ICAM/VCAM coating, electrode arrays were first coated with 4 µg/mL

of AffiniPure F(ab')2 Goat Anti-Human IgG (Jackson ImmunoResearch) for 1h at

37°C in 20mM Tris, pH 8.0; 150mM NaCl; 1mM CaCl2; 2mM MgCl2. BSA (1%) was

added for 30 min at 37°C to block non-specific binding. After blocking, human

recombinant Fc-ICAM-1 or Fc-VCAM-1 were added at a concentration of 1 µg/mL

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and incubated for 16 h at 4°C. The electrical resistance of cells (500,000 cells per

well) seeded on protein-coated arrays was measured in real-time at a frequency of 32

kHz.

Plate adhesion assays

Cells were added to F(ab’)2 / Fc-VCAM-coated wells of flat-bottom 96-wells

Maxisorp plates (100,000 cells per well). Immediately after cell seeding, stimuli were

added to the wells and cells were allowed to adhere for 30 min at 37°C. Subsequently,

non-adherent cells were removed by washing three times with PBS-0.5% BSA.

Adherent cells were fixed with 3.7% formaldehyde, incubated with Hoechst to stain

nuclei and counted on an Axiovert wide-filed microscope (Zeiss).

Adhesion and transendothelial migration under flow

Confluent HUVEC monolayers on FN-coated parallel-plate flow chamber slides (µ-

slide VI0.4, Ibidi) were treated with 10 ng/mL TNF for 16 h. Flow chambers were

mounted on an inverted widefield fluorescence microscope (Axiovert 200, Zeiss)

fitted in a 37°C and 5% CO2 chamber. ShCtrl- and shPrP-U937 cells were differently

labeled with either calcein Green AM or calcein Red-Orange AM (2.5 µM, Life

Technologies). After washing cells were resuspended in assay buffer (20 mM Hepes

(pH 7.4), 132 mM NaCl, 6 mM KCl, 1.2 mM K2HPO4, 1 mM MgSO4, 1 mM CaCl2,

0.1% D-(+)-glucose, 2.5% human albumin). Differently calcein-labeled shCtrl- and

shPrP-U937 cells were mixed at 1:1 ratio to a final total cell concentration of 1x106

cells/mL and subsequently stimulated with 100 ng/mL phorbol-12-myristate-13-

acetate (PMA) or with 100 ng/mL CXCL12 for 10 min. Stimulated cell mixes were

flown over TNF-activated HUVEC monolayers at a flow rate of 0.5 mL/min

corresponding to a shear stress of 0.9 dynes/cm2. Time-lapse bright-field and

fluorescence images were acquired from four random microscope fields for 30-60 min

using an Orca R2 digital CCD camera (Hamamatsu). Quantification of adherent cells

per frame was performed using the ‘analyze particles’ application of ImageJ after

thresholding of the fluorescence video images. Transendothelial migration of U937

cells was assessed in similar experiments as above except for the immobilization of

CXCL12 on the endothelial surface (100 ng/mL, 15 min at 37°C) prior to U937 cell

perfusion. Time-lapse microscopy was performed with a LSM510 META confocal

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microscope (Zeiss). Scoring of transmigrating cells was based on shape changes (from

round to spread) and loss of brightness in the DIC channel.

Cell motility on immobilized Fc-VCAM-1

IBIDI µ-slides were coated with F(ab')2 Goat Anti-Human IgG and Fc-VCAM-1 as

described for ECIS arrays. Subsequently, CXCL12 was absorbed on the dishes by

incubation for 1 h at 37°C with 100 ng/mL CXCL12 in assay buffer. Motility of cells

on Fc-VCAM-1 was recorded from four microscopic fields with a wide-field

microscope as described above. Cell tracking on videos and calculation of

accumulated distance and velocity was performed using Gradientech Tracking Tool

software.

Flow cytometry

Cells were stimulated with CXCL-12 (100 ng/mL) for 15 min at 37°C, cooled on ice

and labeled with SAF32 anti-PrP antibody and anti-1 or anti-2 integrin antibodies

for 1 h at 4°C. Cells were then fixed in 4% paraformaldehyde, blocked in 50 mM

NH4Cl and incubated with Alexa dye-labeled secondary antibodies. Labeled cells

were analyzed by flow cytometry using a FACSCanto flow cytometer (BD

Biosciences). Integrin activation was induced with 100 ng/mL PMA or with 1mM

MnCl2 for 10 min at 37°C in Hank’s Balanced Salt Solution containing 1mM

Ca2+/Mg2+.

Time-course experiments of Fc-VCAM binding to cells was performed according to

(Chan et al., 2003). For Fc-VCAM/ APC- labelled F(ab’)2 binding we followed the

protocol described in (Konstandin et al., 2006). Fluorescence was measured by flow

cytometry using a FACSCanto flow cytometer (BD Biosciences).

Immunofluorescence

For confocal microscopy analysis, antibody-labeled cells as above were resuspended

in mounting medium (Vectashield) containing DAPI, and mounted on a glass slide.

Alternatively, cells were seeded on Fc-VCAM and CXCL12-coated IBIDI µ-slides,

allowed to migrate/polarize for 20 min and fixed with 4% paraformaldehyde in assay

buffer, blocked using 50 mM NH4Cl in PBS and subsequently permeabilized with

PBS-0.1% Triton X-100 for 2 min. After blocking in PBS-1% BSA, cells were

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subsequently incubated with primary antibodies for 30 min at room temperature and

Alexa488-labelled goat anti-mouse antibodies. To stain nuclei, Hoechst was added in

the washing buffer. Fluorescence images were obtained with the LSM510 META

(Zeiss) or with a SP8 (Leica) confocal microscopes.

For colocalization analysis between PrP and 1 integrin, ROIs were selected on

digital pictures and Pearson’s coefficients were determined using the Colocalisation

Test plugin from ImageJ (Costes randomization method). In polarized cells, ROIs

were selected of the capped and non-capped membrane domains. In non-polarized

cells, ROIs were made around the membrane of each cell halves. We considered that

colocalization existed at Pearson’s coefficient values from 0.5 to 1.0.

DNA constructs

Monomeric EGFP (mEGFP) was inserted between the signal peptide (1-22) and

mature PrP (23-253) by ligation independent cloning. A 121 bp fragment containing

the PrP signal peptide was PCR amplified from human PrP cDNA (TrueClone from

Origene) using MFB001(5’-

GCAGGGGCGCAACAGACCCCGGTGCCACCATGGCGAACCTTGGCTGCTGG

ATGC-3’) and MFB002 (5’-

CCACCAGGCCGGCCAGCACCCGGTCCGCAGAGGCCCAGGTCACTCCATG

TG-3’) and inserted in frame in front of mEGFP in XmaI-digested mEGFP-LIC as

described (Bierings et al., 2012), resulting in sp-mEGFP-LIC. A 754 bp fragment

containing mature PrP was PCR amplified using MFB003 (5’-

GGGCGCGCCTGGTGGGGCCAAGAAG CGCCCGAAGCCTGGAGGATG -3’)

and MFB004 (5’- GCGGCCGCCTGCTCGTCCATCATC

CCACTATCAGGAAGATGAGG -3’) and inserted in frame behind mEGFP in SacII-

digested sp-mEGFP-LIC using ligation independent cloning. All plasmids were

verified by sequence analysis.

Rescue of PrP expression

U937 cells stably expressing shRNA targeting human PrP were cultured for 48 hours

in puromycin-free medium. After 48 hours cells were transfected by electroporation

(Neon Transfection System, Life Technologies) with a GFP-PrP or GFP construct

according to manufacturers’ recommendations. GFP-positive cells were sorted using a

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FACSAria II cell sorter (BD biosciences). Cells were allowed to recover for 2 hours

in complete medium before seeding on Fc-VCAM-coated IBIDI µslides.

GTPase activation assay

Cells were lysed in 25 mM Tris-HCl, pH 7.2, 150 mM NaCl, 10 mM MgCl2,

1% NP-40 and 5% glycerol and cOmplete Protease Inhibitor Cocktail Tablets, Roche.

RhoA-GTP was isolated from cell lysates using a pull-down assay as previously

described (Alblas et al., 2001).

Immunobloting

Cells were stimulated in suspension and lysed in cold NP-40 buffer (25 mM Tris-HCl,

pH 7.2, 150 mM NaCl, 10 mM MgCl2, 10 mM CaCl2 1% NP-40 and 5% glycerol)

containing Halttm protease and phosphatase inhibitor cocktail (Thermo Scientific).

Phospho-ERM and total moesin levels were determined by Western blotting. Pixel

density was determined using ImageJ. Data represent phospho-ERM/moesin ratios

and were normalized to t=0 of shCtrl cells.

To determine phospho-cofilin and total cofilin levels, U937 cells were lysed in sample

buffer after 30 min adhesion to VCAM-coated plates. Equal loading was controlled

by detection of total cofilin. Pixel density values was determined using ImageJ. Data

represent phospho-cofilin/cofilin ratios. Values were normalized to shCtrl.

Actin polymerization assay

CXCL12-induced changes in intracellular content of F-actin were measured by

Alexa488-phaloidin staining as described in (Voermans et al., 2001). Labeled cells

were analyzed by flow cytometry using a FACSCanto flow cytometer (BD

Biosciences)

Statistical analysis

Data are represented as mean±SD or mean±SEM. Statistical comparisons of means

were performed using either two-tailed Student t test, one-way ANOVA with

Bonferroni post hoc test or two-way ANOVA (Bonferroni’s multiple comparisons

test)post hoc test. Statistical significance is represented by asterisks: * p<0.05;

**p<0.01; ***p<0.001.

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Acknowledgements

We are grateful to Dr Peter Hordijk for critically reading our manuscript. We also

thank Erik Mul for his assistance with flow cytometry assays and Nicoletta Sorvillo

for helping with monocyte isolation. We are grateful to Maaike Scheenstra for sharing

with us her protocol for monocyte transduction and to Dr Keith Burridge and Dr

Metello Innocenti for their helpful comments.

This work was financed with a grant from Sanquin Blood Supply (PPOC 11-006). RB

was supported by a European Hematology Association Research Fellowship.

Author contribution

D.D.R., S.T., E.V-E., C.P. and R.B. performed the research and analyzed the data;

M.F-B. designed the research, analyzed the data and wrote the paper. D.G.

contributed reagents.

D.R. and S.T. contributed equally to this study.

Conflict of interest disclosure: the authors declare no competing financial interests.

Correspondence: Mar Fernandez-Borja, Department of Molecular Cell Biology,

Sanquin Research, Plesmanlaan 125, 1066CX Amsterdam, The Netherlands; e-mail:

m.fernandez@sanquin.nl

Competing interests

The author(s) declare no competing financial interests

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Figures

Figure 1. PrP resides in cholesterol-rich membrane domains on the surface of

human pro-monocytic U937 cells. (A) Detection of PrP in live cells by flow

cytometry using the SAF32 antibody. Mouse IgG2b was used as isotype control. (B)

Triton X-100 cell lysates were fractionated in Optiprep density gradients. Fractions

were collected from the top, proteins precipitated with trichloroacetic acid and

analyzed by immunoblotting for the presence of PrP, the lipid raft-marker flotillin-1

and the non-lipid raft marker transferrin receptor (TfR). PrP and flotillin-1 were

enriched in the top fractions consistent with their localization in lipid rafts. (C) U937

cells were stimulated or not with 100 ng/mL CXCL12 for 10 min at 37°C and then

cooled on ice for PrP and integrin detection with SAF32 (PrP) and P5D4 (β1 integrin

subunit). Pearson’s colocalization coefficients between PrP and 1 integrin were

determined in the selected ROI’s (A and B) depicted in the scheme, both in polarized

and non-polarized cells. In polarized cells, A included the integrin cluster and B, the

remaining membrane. In non-polarized cells, each, A and B, correspond to one half of

the membrane. The A/B ratio indicates the relative colocalization measured in A with

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respect to B. Scale bar, 10 µm. (D) Stable U937 cell lines were generated by

puromycin-selection of cells transduced with lentiviruses expressing small hairpin

RNAs to human PrP (shPrP-1, -2 and -3) or with non-targeting shRNA (shCtrl). PrP

expression was examined by flow cytometry using the SAF32 anti-PrP Ab. Graphs

represent the average of three independent transduction/ selection experiments.

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Figure 2. PrP regulates U937 cell and primary monocyte adhesionto 1 integrin

ligands. (A) CXCL12-induced migration of shCtrl- and shPrP-U937 cells was

determined using a Transwell assay. Due to the inter-experimental variation of

absolute migration values, the migration values of shCtrl-U937 were set to 100% to

determine statistical significance of the differences. Data are from three independent

experiments performed in triplicate. Statistical analysis was performed using 2-tailed

unpaired Student t-test. (B) CXCL12-induced migration of U937 cells incubated with

either anti-PrP SAF61 antibody or IgG2a was determined using a Transwell assay as

in (A). (C) Representative ECIS experiment showing real-time changes in electrical

resistance upon adhesion of shCtrl- and shPrP-U937 cells to FN or BSA. The black

arrow indicates addition of cells to the electrodes and the green arrow indicates PMA

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addition to stimulate cell adhesion. (D) PMA-stimulated adhesion of shCtrl- and

shPrP-U937 to BSA and FN was determined dividing the resistance at t=1h by the

resistance at t=0 after PMA addition (n=3). Statistical significance was analyzed using

two-way ANOVA); (E) PMA-stimulated shCtrl- and shPrP-U937 cell adhesion to

VCAM-1 and ICAM-1 was calculated as in (D) (n=6 for VCAM; n=4 for ICAM).

Statistical significance was analyzed using two-way ANOVA. (F) ECIS adhesion

assay of shCtrl- and shPrP-primary human monocytes as in C. (G) Quantification of

PMA-stimulated adhesion as in (D) (n=3). Statistical significance was analyzed using

unpaired two-tailed Student t-test.

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Figure 3. PrP regulates U937 cell and primary monocyte adhesion to endothelial

cells under shear stress. (A) Mixed populations of red-labelled shCtrl- and green-

labelled were incubated with PMA or CXCL12 for 10 min before being perfused over

TNF-stimulated endothelial monolayers. Upper panels show DIC and fluorescence

snapshots of 30 min video recordings of U937 adhesion to the endothelium (arrow

indicates the direction of flow). Lower panels show the result of image processing

with ImageJ software to automatically count adherent U937 cells. Bar: 50 µm. (B)

Graph shows the total number of CXCL12-stimulated cells adherent to the

endothelium in three microscopic fields of a representative flow experiment. (C)

Same as in (B) for PMA-stimulated cells. (D) Percentage of CXCL12-stimulated

shCtrl- and shPrP-U937 cells adherent to the endothelium per microscopic field after

10 min of flow (n=3). (E) Same as in (D) for PMA-stimulated cells. (F) Same as in

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(D) for CXCL12-stimulated primary monocytes. Statistical analysis was performed

using unpaired two-tailed Student t-test.

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Figure 4. Defective uropod formation in migrating PrP-silenced cells. (A) shCtrl

and shPrP-U937 cells were seeded on Fc-VCAM-1/ CXCL12-coated plates and

recorded using a widefield microscope. Figure shows snapshots of these video

recordings. Migrating shCtrl-U937 cells develop long uropods (arrowheads). In

contrast, uropod formation is impaired in shPrP-U937 cells. Bar graphs represent the

quantification of accumulated distance, cell velocity and directness by manual cell

tracking using Gradientech Tracking Tool software. Statistical analysis was

performed using two-tailed unpaired Student t-test. (B) shCtrl- and shPrP-U937 cells

were seeded on Fc-VCAM-1 and CXCL12-coated IBIDI -slides, allowed to migrate

for 20 min, and processed for immunofluorescence detection of phospho-ERM

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proteins and 1 integrins. The bar graph shows the average length of the uropod in

cells migrating on VCAM as described above (n=3, ≥40 cells (each for shCtrl- and

shPrP-U937 cells) per experiment). Statistical analysis was performed using unpaired

two-tailed Student’s t-test. Bar: 20 m (C) shPrP-U937 cells were transfected with

GFP-PrP or GFP (negative control) cDNA by electroporation and sorted by flow

cytometry to enrich for transfected cells. Sorted cells were seeded on immobilized Fc-

VCAM-1/ CXCL12 and fixed after for 20 min for the immunodetection of 1

integrins and PrP. The bar graph shows average uropod length in GFP-PrP and GFP-

expressing cells (n=3). Statistical analysis was performed using unpaired two-tailed

Student’s t-test. Bar: 10 m.

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Figure 5. PrP silencing impairs ERM protein activation. (A) Phosphorylated ERM

proteins were detected with an antibody that recognizes a phosphorylated threonine

residue conserved in all three proteins. Blots were reprobed with antibodies against

total moesin. Quantification of band density was performed as described in the

methods section (n=5). Statistical analysis was done using two-way ANOVA

(Bonferroni’s multiple comparisons test). Since this test takes into account all the time

points, differences at t=0 of CXCL12 stimulation, although with similar values as t=0

for PMA stimulation, is not significant when compared with later time points. (B)

CD44 and flotillin-1 were immunodetected in shCtrl- and shPrP-U937 cells migrating

on immobilized VCAM. Bar: 10 m.

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Figure 6. Increased motility and membrane protrusive activity of PrP-deficient

U937 cells on the endothelium under shear stress. (A) DIC and fluorescence

snapshots of a video recording of PMA-stimulated calcein Red-Orange-labeled

shCtrl-U937 cells and calcein Green-labeled shPrP-U937 cells on TNF-activated

endothelial cells under shear stress. ShPrP-U937 cells undergo shape changes while

crawling over the endothelium (lower green cell) while the shCtrl-U937 cell is

motionless and maintains a spherical shape. Time is indicated on the figures in

min:sec. (B) Percentage of shCtrl- and shPrP-U937 cells that displayed dynamic

membrane protrusions after adhesion to endothelial cells under flow (n=3). (C)

Percentage of adherent shCtrl- and shPrP-U937 cells that migrated across the

endothelium (n=3). Statistical analysis was performed using unpaired two-tailed

Student’s t-test.

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Figure 7. PrP regulates integrin avidity. (A) Flow cytometry analysis of total and

activated 1 integrins on shCtrl- (black line) and shPrP- (red line) U937 cells treated

or not with CXCL12, PMA or Mn2+ for 10 min at 37°C. Cells were incubated with

mouse IgG1 as isotype control (grey shaded area). No differences for total or

activated integrins were detected between shCtrl- and shPrP-U937. (B) Cells were

stimulated for different time-points with CXCL12, PMA or Mn2+ in the presence of

Fc-VCAM. Cell-bound Fc-VCAM was detected with APC-labelled F(ab’)2 goat anti

human IgG and analyzed by flow cytometry (n=3). (C) Similar experiment as in (B)

except that cells were mixed with pre-formed multimeric Fc-VCAM/ F(ab’)2-APC

complexes (n=3). Statistical analysis was performed using two-way ANOVA.

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Figure 8. PrP silencing enhances RhoA activation. (A) A pull-down assay was used

to assess the levels of GTP-bound active RhoA in shCtrl- and shPrP-U937 stimulated

with CXCL12 or PMA for the indicated times (in min). Total RhoA was detected in

the corresponding whole cell lysates for input. Graphs represent quantification of the

bands as described in the methods section (n=6). (B) Cell F-actin content was

determined at different time points of CXCL12 stimulation by flow cytometry using

Alexa488-labelled phaloidin (n=3). (C) Cells were incubated for 30 min at 37°C with

1 or 5 m cytochalasin B and subsequently seeded on electrode arrays for the ECIS

adhesion assay. Figure shows a representative graph corresponding to one of three

independent experiments showing changes in adhesion from the moment of cell

addition. Bar graph represents the increase in resistance from the moment of cell

addition until 1h after PMA addition (n=3). (D) Phospho-cofilin and total cofilin were

detected by immunoblotting of cell lysates of cells after 30 min adhesion to

immobilized Fc-VCAM. Bar graphs show the quantification of band densities of three

independent experiments. Statistical analysis was performed using two-way ANOVA

(A, B, C) and two-tailed Student’s t-test (D).

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