Correspondence to Bernhard Wehrle-Haller: Bernhard.Wehrle-Haller{at}medecine.unige.ch
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C. Cluzel's present address is Institut de Biologie et Chimie des Proteines, Lyon Cedex 07, France.
F. Saltel, F. Paulhe, and B. Wehrle-Haller's present address is Dept. of Cellular Physiology and Metabolism, Centre Medical Universitaire, 1211 Geneva 4, Switzerland.
Abbreviations used in this paper: cRGD, cyclic RGD; cD, cytochalasin D; IRM, interference reflection microscopy; PI(4,5)P2, phosphoinositol-4,5-bisphosphate; TIRF, total internal reflection fluorescence; WT, wild type.
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Introduction |
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Integrins are noncovalently linked heterodimeric receptors that exist in low- and high-affinity states (Hynes, 2002). In the low-affinity state, the ectodomains of the and ß subunits are in a folded configuration with laterally associated transmembrane and cytoplasmic domains. Upon integrin activation, the distal parts of both ectodomains swing open in a switchblade motion, followed by the separation of the transmembrane and cytoplasmic domains (Xiao et al., 2004). This allosteric switch, or "integrin activation," can be induced by effector binding to the cytoplasmic tail of the
and ß subunit (Vinogradova et al., 2002; Katagiri et al., 2003; Tadokoro et al., 2003) or by ligand binding to the ectodomain (Takagi et al., 2002). Once the integrin heterodimer is in its open configuration, the cytoplasmic domain of the ß integrin chain is linked via cytoplasmic adaptor proteins, such as talin, to the actin cytoskeleton (Liddington and Ginsberg, 2002; Critchley, 2004).
The molecular mechanisms that control the lateral assembly of integrins ("clustering") to form focal adhesions are controversial. It has been proposed that the process of integrin clustering is intimately linked to the switch in its affinity state. In vitro studies have suggested that the activation-induced separation of the integrin transmembrane domains can induce integrin clustering by the respective homooligomerization of the and ß transmembrane domains (Li et al., 2003). However, this model has been contested based on disulfide bond scanning of the exofacial portions of the transmembrane domains of activated integrins expressed in living cells and extensive mutagenesis of the transmembrane domains (Luo et al., 2004; Partridge et al., 2005). Therefore, it remains an open question whether (and how) integrin activation is coupled to integrin clustering and focal contact remodeling in living cells. We analyzed this issue by expressing wild-type (WT) and constitutively activated EGFP-tagged
vß3 integrins within living cells and investigating the requirement of extracellular ligand binding, recruitment of cytoplasmic adaptors, and membrane lipid composition for the clustering of activated integrins. In addition, we determined by FRAP how constitutive activation of integrins alters their dynamic remodeling within focal contacts.
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Results |
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In adherent cells, WT ß3-EGFP integrins formed brightly fluorescent clusters at the cell periphery, representing focal contacts (Ballestrem et al., 2001; Fig. 1 A). In addition, nonclustered integrins were found evenly dispersed within the plasma membrane (Fig. 1 A). Because the population of cell surface integrins is in equilibrium between the activated and nonactivated forms, it is not clear to what extent activated integrins are present in the clustered or nonclustered state. To shift the equilibrium toward activated states of integrins, we treated cells with Mn2+. Only minutes after Mn2+ activation, we observed the formation of numerous irregularly shaped integrin clusters underneath the main cell body, in addition to the preexisting clusters of integrins at the periphery of the cell (Fig. 1 B). Similar to Mn2+-activated WT ß3 integrin, the activated ß3 integrin mutants D723A and N305T formed clusters underneath the entire cell body and in the periphery of the cell (Fig. 1, C and D). In contrast, the inactive ß3 integrin mutant (D119Y), lacking the ability to bind ligand, was found evenly dispersed at the cell surface, exhibiting no apparent integrin clustering (Fig. 1 E). These results demonstrate that integrin activation correlates with integrin clustering and suggest a role for ligand binding in this process (Miyamoto et al., 1995).
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Time course of integrin clustering
The presence of clusters of activated integrins under the main cell body is unusual. Because focal contacts and the proximity of the cell membrane to the substrate can be revealed by interference reflection microscopy (IRM), we analyzed the time course of Mn2+-induced integrin clustering in respect to the IRM image. Before Mn2+ stimulation, ß3 integrins were clustered within peripherally located focal contacts, appearing dark in corresponding IRM images (Fig. 1, G and J; Verschueren, 1985). Streaklike IRM-dark structures located toward the center of the cell did not exhibit ß3 integrin clustering (Fig. 1, G and J, insets). Similarly, dotlike close contacts located underneath the main cell body were not positive for ß3 integrins. Shortly after Mn2+ activation, the fluorescence intensity of preexisting peripheral focal contacts increased, and previously ß3-negative, streaklike dark contacts appeared ß3 positive (Fig. 1, H and K). Subsequently, dotlike close contacts located underneath the main cell body appeared ß3 integrin positive, although with lower fluorescence intensity compared with peripheral focal contacts (Fig. 1, I and L). Hence, de novoactivated integrins gradually appear in streak- and dotlike dark and close contacts underneath the main cell body.
Specific talin recruitment to F-actinindependent integrin clusters
Clusters of Mn2+ or mutational activated integrins were found underneath the main cell body, a cellular localization rarely populated by focal contacts. Because integrins clustered within focal adhesions are mechanically linked to the actin cytoskeleton, we analyzed whether actin and/or focal adhesion adaptor proteins are recruited to de novoformed integrin clusters. We analyzed whether phalloidin reactive F-actin was localized to Mn2+-induced integrin clusters. Mn2+ activation resulted in the formation of integrin clusters in cellular regions devoid of F-actin (Fig. 2, A and A', inset). De novo formation of integrin clustering independent of F-actin was confirmed by treatment of spread cells with cytochalasin D (cD) before Mn2+ activation. Despite the destruction of the actin cytoskeleton by cD, Mn2+-activated ß3 integrins formed clusters (Fig. 2, B and B'). Because actin fibers were dispensable for integrin clustering, we asked whether adaptor proteins were recruited to clusters of activated integrins. Immunofluorescence staining with antitalin antibodies revealed an overlap with all clustered EGFP integrins in Mn2+-treated cells (Fig. 2, C and C'). This confirms the data that talinintegrin association is important for the formation of focal adhesions (Priddle et al., 1998). In contrast to talin, the focal adhesion adaptor proteins vinculin, paxillin, and FAK, as well as antiphosphotyrosine antibodies, did not associate with de novoformed clusters of activated integrins (Fig. 2, DG). An identical result was obtained in ß3 integrin negative CS-1 hamster melanoma cells that had been transfected with non-EGFPtagged ß3 integrins (Fig. 2, H and I). This suggests that the observed clustering of activated integrins and selective talin recruitment is not influenced by the EGFP tag.
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Integrin clustering requires PI(4,5)P2
It has been demonstrated that peptides, representing the cytoplasmic tail of integrins, bind to talin. In addition, phosphoinositol-4,5-bisphosphate (PI[4,5]P2) binding to talin induces a conformational change that facilitates the interaction of the talin head domain with the cytoplasmic tail of ß3 integrins (Martel et al., 2001). Therefore, we asked whether PI(4,5)P2 is a critical cofactor in the clustering of high-affinity integrins. We treated ß3 integrinexpressing cells with increasing doses of neomycin sulfate, a drug known to selectively bind and sequester PI(4,5)P2 (Arbuzova et al., 2000; Laux et al., 2000). The sequestration of PI(4,5)P2 by 10 mM neomycin dramatically reduced the formation of ß3 integrin clusters in the periphery of control cells (Fig. 3 A), as well as Mn2+-induced, de novo integrin clusters underneath the main cell body (Fig. 3 B). Despite the reduction of integrin clustering under the main cell body, some remaining integrin clusters were found in the cell periphery often associated with filopodia. These thin, fingerlike integrin clusters were in contact with the substrate as suggested by the corresponding IRM image (Fig. 3, A and B, insets). Similar to Mn2+-stimulated WT cells, 10 mM neomycin suppressed peripheral focal contacts and integrin clusters under the main cell body in D723A and N305T mutant ß3 integrintransfected cells (unpublished data). To better characterize the effect of neomycin on integrin clustering, we performed dose response analysis with WT ß3 integrinexpressing cells. Mn2+-induced integrin clustering was efficiently prevented at 1 mM neomycin sulfate (Fig. 3 C). To exclude the possibility that the neomycin sulfatedependent inhibition of integrin clustering was indirectly caused by the drug's effect on the actin cytoskeleton (Laux et al., 2000; Kwik et al., 2003), we treated cells with cD before Mn2+ stimulation. Irrespective of the state of the actin cytoskeleton, neomycin sulfate prevented the formation of Mn2+-induced integrin clusters (unpublished data). To test whether PI(4,5)P2 was also important for the maintenance of integrin clusters, we tested whether its sequestration would affect previously clustered integrins. The addition of 10 mM neomycin sulfate to Mn2+-stimulated cells dispersed integrin clusters within 1 h (Fig. S4, available at http://www.jcb.org/cgi/content/full/jcb.200503017/DC1). Nevertheless, the dispersed but activated integrins were still able to link the cells to the substrate as revealed by IRM (unpublished data). These results demonstrate that PI(4,5)P2 is involved in the induction and stabilization of the lateral association of integrins, either by increasing the affinity of talin for integrins (Martel et al., 2001) or by oligomerization of the integrintalin complex through interaction with lipid domains containing PI(4,5)P2.
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To determine whether integrin clustering requires "immobilized" in contrast to "soluble" extracellular ligands, we performed Mn2+-induced integrin stimulation in the presence of increasing doses of cyclic RGD (cRGD) peptides. Importantly, we chose a concentration of cRGD that did not completely inhibit the recruitment of vß3 integrins into peripheral focal adhesions (Fig. 3 G). At concentrations of 10 µM cRGD, which inhibited binding of soluble
vß3 integrin to immobilized ligand in an ELISA-type assay (Legler et al., 2001), the formation of Mn2+-induced de novo integrin clusters underneath the main cell body was completely blocked (Fig. 3 I). These data demonstrate that both the cytoplasmic and the extracellular ligand-binding domains are essential for the clustering of
vß3 integrins. Moreover, immobilized extracellular ligands are required to stabilize nascent clusters of activated integrins at the cell surface.
The head domain of talin induces integrin clustering
As demonstrated in Fig. 2, the specific recruitment of talin to Mn2+-induced integrin clusters suggests that talin is critically involved in the formation of integrin clusters. Moreover, because talin exists as an anti-parallel dimer with the integrin-binding head domains positioned at both ends (Isenberg and Goldmann, 1998), talin represents a bona fide intracellular integrin cross-linker. To test whether the dimeric form of talin is required for integrin clustering, we overexpressed the monomeric integrin binding head domain of talin (residues 1435; Yan et al., 2001) as a CFP-tagged chimera in stable ß3-GFP integrinexpressing cells, with the intention to dominantly suppress integrin clustering. Instead, the overexpression of the isolated head domain of talin induced an increase in integrin clustering (Fig. 4, AC). Similarly, Mn2+ stimulation of cells that overexpressed the head domain of talin induced a greater increase in integrin clustering, covering more than half of the cell surface (Fig. 4, D and E). In most cells, the cytoplasmic expression of talin was very high, making it impossible to determine whether the talin head domain only activated integrins or was also involved in their clustering. However, after Mn2+ stimulation, high levels of the talin head domain were no longer required for integrin activation, which nevertheless resulted in efficient clustering of integrin at low talin expression levels (Fig. 4, G and H). In these clusters (Fig. 4 G), we detected a colocalization with the head domain of talin (Fig. 4 H). These data suggest that the monomeric talin head domain possesses the ability to induce integrin activation and clustering. Because talin-dependent integrin clustering is sensitive to neomycin treatment (unpublished data), integrin clustering may be induced by the binding of the talin head domain to multivalent PI(4,5)P2 lipid domains.
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Discussion |
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The driving force of lateral integrin association
The nature of the driving force that induces the lateral association of activated integrins is a matter of debate. One possibility put forward by Li et al. (2003) is that homophilic interactions between the transmembrane domains of activated integrins drive lateral clustering. This model implies that integrin clustering can occur independently of other integrin-binding proteins as long as the transmembrane domains of the and ß integrin subunit are physically separated from each other (e.g., separation induced by integrin activation). Other results, however, do not support the view that activated integrins can cluster spontaneously. Electron microscopic images of purified, activated
IIbß3 integrins that are incorporated into lipid vesicles or planar membranes give no evidence for spontaneous clustering of activated integrins (Erb et al., 1997). In addition, Mn2+-activated
vß3 integrins do not cluster spontaneously when exposed to a laminin-1 substrate to which they are unable to bind (unpublished data). Moreover, the disulfide bond scanning of the exofacial portions of the integrin
IIb and ß3 transmembrane domains did not reveal a specific interaction of these domains after integrin activation in living cells (Luo et al., 2004). We now demonstrate that activated
vß3 integrins require immobilized substrate, PI(4,5)P2 lipids, and the focal adhesion adaptor protein talin for clustering. Our data suggest that integrin clustering is controlled by a simple associationdissociation reaction that is influenced by the density of activated integrins in the plasma membrane. However, the equilibrium of this reaction can be shifted depending on the availability of immobilized substrate, PI(4,5)P2, and talin (Fig. 6 A).
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In addition to talin cleavage, the reversible phosphorylation of PI(4)P may be a way to control focal adhesion assembly or disassembly (Fig. 6 B). Signal-dependent recruitment of PIPKI to the talin head domain could create a local focal adhesionassociated source of PI(4,5)P2 that in turn would stimulate talin recruitment and integrin activation (Di Paolo et al., 2002; Ling et al., 2002; Barsukov et al., 2003). Thus, PIPKI
activity may represent the driving force for integrin remodeling in sliding focal adhesions at the cell rear.
Independent of the "intracellular" regulation of integrin dynamics in focal contacts, our results demonstrate that the rate-limiting step for the assembly and disassembly of integrin clusters is the time during which the activated integrin is bound to immobilized extracellular ligands. This is supported by the slow integrin exchange dynamics of the N305T ectodomain mutant, suggesting that the flexibility, and hence the extracellular ligand affinity, of the ectodomain controls integrin cluster dynamics.
Integrin clusters are precursors of focal adhesions
De novo Mn2+-induced ß3 integrin clusters specifically recruit talin but not other adaptor proteins such as vinculin, paxillin, or FAK. This is in contrast to the recruitment of FAK and other signaling components, but not talin, to antibody cross-linked RGD peptide-activated clusters of 5ß1 integrin (Miyamoto et al., 1995). It is currently not known which mechanisms are responsible for these different patterns of integrin adaptor recruitment. In the case of
vß3 integrin however, we propose that the integrinPI(4,5)P2talin complex represents an intermediary step during the assembly of focal adhesions, which are rendered visible by the forced activation of large numbers of integrins. Accordingly, when cells are treated for a prolonged time with Mn2+ (e.g., 2 h), an increasing fraction of integrin clusters can be found to associate with vinculin and F-actin (unpublished data). Consequently, the integrinPI(4,5)P2talin complex should be found in locations where focal complexes normally assemble, such as in the lamellipodium. In fact, comparable talin- and integrin-containing immature focal complexes have been found at the leading edge of advancing lamellipodia (DePasquale and Izzard, 1991; Lee and Jacobson, 1997). These structures further mature by recruiting vinculin,
-actinin, and FAK (DePasquale and Izzard, 1991; Lee and Jacobson, 1997). This evolution suggests that talin-associated activated integrins are the progenitors of focal complexes within the lamellipodium. This view is supported by a recent report demonstrating the recruitment of talin into ß3 integrin containing focal complexes before the appearance of vinculin or FAK (Zaidel-Bar et al., 2003). As an exception to the rule, paxillin is present in focal complexes of migrating endothelial cells at the same time as ß3 integrins (Zaidel-Bar et al., 2003). The difference to our data could be explained by the specific recruitment of paxillin to
4 integrins that may cluster together with
vß3 integrin at the leading edge of endothelial cells (Goldfinger et al., 2003).
We propose, therefore, that the formation of focal complexes within a lamellipodium includes several steps: PI(4,5)P2 is synthesized in the lamellipodium in a Rac1- and ARF6-dependent manner (Honda et al., 1999). PI(4,5)P2 induces a conformational change in talin that associates and activates ß3 integrins (Martel et al., 2001; Calderwood et al., 2002), forming "preadhesion complexes." Simultaneously, PI(4,5)P2 stimulates vinculin and ezrinradixinmoesin family proteins to bind to talin and to link the actin cytoskeleton to the plasma membrane, respectively (Yin and Janmey, 2003), which is a process facilitated by the transient interaction of vinculin with the actin nucleation factor Arp2/3 (DeMali et al., 2002). Therefore, a focal complex forms in the lamellipodium as a result of the intersection of several intra- and extracellular systems requiring immobilized ligand, integrin activation, actin polymerization, and PI(4,5)P2 synthesis.
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Materials and methods |
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Cell culture, transfection, and inhibitor treatment
Mouse B16F1 melanoma cells and hamster CS-1 melanoma cells were grown in DME containing 10% FCS, glutamine, and antibiotics (Thomas et al., 1993; Ballestrem et al., 2001). Cells were transfected using Fugen 6 (Roche) and selected for stably expressing cells in the presence of 1.2 mg/ml B16F1 or 0.6 mg/ml CS-1 G418 (Becton Dickinson). Stably transfected B16F1 cells were FACS sorted, and clones were selected for their expression of EGFP fluorescence and cell surfaceexposed vß3 integrins using the Kistrin-CD31 fusion protein SKI-7 and the rat anti-CD31 mAb GC51, as previously described (Ballestrem et al., 2001; Legler et al., 2001) or using hamster antimouse ß3 integrin in the case of the D119Y mutant (2C9.G2; BD Biosciences). Nontagged ß3 integrintransfected CS-1 cells were selected for their SKI-7 reactivity and their capacity to adhere to tissue culture dishes. Neomycin sulfate, cD (both from Sigma-Aldrich), and cRGD peptides (cRGD; Bachem; Legler et al., 2001) were prepared as stock solutions in PBS. Inhibitor studies were all performed in complete medium.
Measurement of integrin clustering
WT and mutant B16F1 ß3-EGFPexpressing cells were cultured overnight in complete medium in glass bottom dishes, fixed for 10 min with 4% PFA, and rinsed with PBS. Mn2+ activation (0.5 µM Mn2+) of B16F1 or CS-1 cells was performed for 20 min in complete or inhibitor-containing complete medium. EGFP fluorescence and respective IRM images were acquired with identical excitation and exposure settings using a Plan Neofluar 63x NA 1.4 oil-immersion objective on an inverted confocal microscope (model LSM510; Carl Zeiss MicroImaging, Inc.), focusing on the glass coverslip by using the maximal reflection of the laser light. Intensity histograms of cells were obtained after smoothing (3 x 3 kernel), background subtraction, and manual selection of the cell surface using MetaMorph software (Molecular Devices) and exported to Excel (Microsoft) for further analysis. Histograms were normalized in respect to the cell surface area and aligned according to the peak of the histograms corresponding to the membrane fluorescence of the cell before averaging (n > 20). Cumulative intensity histograms were obtained by multiplication of the number of pixels (in percent) with the respective gray value (Fig. S1).
The relative area of integrin clusters in respect to the entire cell surface was obtained from total internal reflection fluorescence (TIRF) images of at least 20 cells per condition. TIRF images of ß3-EGFP integrin fluorescence were obtained with a Plan Neo-Fluar 100x NA 1.45 objective mounted on an Axiovert 100M (both from Carl Zeiss MicroImaging, Inc.) equipped with a 12-bit digital charge-coupled device camera (model Orca 474295; Hamamatsu Photonics) controlled by the Openlab software (Improvision). Intraobjective TIRF was obtained with a 488-nm laser through a TIRF adaptor (TILL Photonics). Image analysis was performed after manually setting of the intensity threshold using MetaMorph software.
Measurement of integrin intensities
Relative integrin fluorescence intensities in control and Mn2+- or neomycin sulfatetreated cells were essentially measured as described in Ballestrem et al. (2001). In brief, 10-bit-wide field fluorescent images were acquired with identical settings on an Axiovert 100TV using Openlab software. For each image, the fluorescence intensity was determined from triplicate measures of the background outside the cell, inside bright peripheral contacts, in the cell membrane, and, when appropriate, in Mn2+-induced integrin clusters. The respective integrin fluorescence was obtained by deduction of the background.
Antibodies, immunofluorescence, and the digital processing of images
Cells grown overnight in complete medium were fixed with 4% PFA in PBS for 10 min, permeabilized, and blocked in 0.1% Triton X-100 and 1% BSA in PBS for 20 min. Mouse mAbs to vinculin, talin recognizing the rod domain of talin (both from Sigma-Aldrich), P-Tyr (4G10; Upstate Biotechnology), FAK (F15020), and paxillin (P13520; both from BD Transduction Laboratories) were applied in 1% BSA-PBS for 1 h. After being washed in blocking solution, Texasred conjugated goat antimouse antibodies (Jackson ImmunoResearch Laboratories) were applied for 1 h and subsequently washed as above. Preparations were stored in PBS and images were collected at RT using a Ph3 Plan-Apochromat 63x NA 1.40 objective on an Axiovert 100TV microscope equipped with a 10-bit digital charge-coupled device camera controlled by Openlab software. Confocal images were obtained as described in Measurement of integrin clustering on a confocal microscope. Due to technical reasons, confocal images of EGFP-integrin and ECFP-talin double-transfected cells were acquired individually after changing the dichroic mirror. For publication, a Gaussian blur (0.4) was applied to all confocal images. In addition, background and contrast were adjusted using the adjust Level command in Photoshop (Adobe).
Substrate patterns
Substrate patterns of 5 x 5 µm2 squares were prepared using a variation of the microcontact printing technique (Csucs et al., 2003). Polydimethylsiloxane stamps carrying the corresponding pattern (Csucs et al., 2003) were incubated for 1 h with a mixture of 20 µg/ml of Alexa Fluor 568labeled fibronectin and 10 µg/ml of nonlabeled vitronectin in PBS. Inked stamps were placed onto glass coverslips for 30 s to ensure contact between the stamp and the glass surface. After rinsing the coverslip with PBS, noncoated areas of the coverslip were blocked for 20 min in 1 mg/ml of polylysine-g-polyethyleneglycol copolymer in 10 mM Hepes, pH 7.4, which is proven to be efficient in repelling serum-protein absorption to the glass coverslip (Huang et al., 2001; VandeVondele et al., 2003). Vitronectin and fibronectin were obtained from Sigma-Aldrich and fibronectin was labeled with an Alexa Fluor 568 labeling kit as recommended by the manufacturer (Molecular Probes). Cells were plated overnight in BSA containing medium and were shifted to 3% FCS containing medium 1 h before imaging.
FRAP
B16F1 cells were cultured in glass bottom dishes for 24 h in complete culture medium. Medium was replaced by F12 medium containing glutamine, antibiotics and 10% FCS before FRAP analysis. FRAP was performed at 37°C on an LSM510 inverted microscope equipped with a heated stage and CO2 control essentially as described in Ballestrem et al. (2001). To reduce loss of fluorescence due to bleaching during the recovery period, we reduced the laser power of the 488-nm line to 0.5% at a maximal laser output of 50%. To improve light collection, the pin hole was partially opened (to 200 µm). To ensure maintenance of the focus during the recovery period, the IRM was recorded simultaneously.
Online supplemental material
Fig. S1 further explains the procedure used to evaluate integrin clustering from confocal images. In addition, the phenotype of integrin "shedding" observed in the N305T ß3 integrin mutant transfected cells is illustrated in Fig. S2. Fig. S3 demonstrates the ability of the cytoplasmic tail of ß1 integrin to mediate Mn2+-induced integrin clustering. In Fig. S4 the role of PI(4,5)P2 in maintaining Mn2+-induced integrin clusters is shown by their dispersal in response to neomycin sulfate treatment. Videos 16 represent the corresponding FRAP sequences of the cells from which the boxed areas are shown in Fig. 5 (AG). Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200503017/DC1.
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Acknowledgments |
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C. Cluzel and F. Saltel were supported by grants for post-doctoral fellowships of the French Foundation for Medical Research and the French Association for Cancer Research. The work was supported by research grants from the Swiss Cancer Ligue (KFS 412-1-1997) and the Swiss Science Foundation to B.A. Imhof and B. Wehrle-Haller (31-059173.99, 31-64000.00, and 3100A0-103805).
Submitted: 3 March 2005
Accepted: 17 September 2005
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