Osmotic stress-induced remodeling of the cortical cytoskeleton

Caterina Di Ciano1, Zilin Nie2, Katalin Szászi3, Alison Lewis1, Takehito Uruno4, Xi Zhan4, Ori D. Rotstein1, Alan Mak2, and András Kapus1

1 Department of Surgery, Toronto General Hospital, University Health Network and University of Toronto, M5G 1L7; 2 Department of Biochemistry, Queen's University, Kingston, K7L 3N6; 3 Division of Cell Biology, Hospital for Sick Children, Toronto, Ontario, M5G 1X8 Canada; and 4 Department of Experimental Pathology, Holland Laboratory, American Red Cross, Rockville, Maryland 20855


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Osmotic stress is known to affect the cytoskeleton; however, this adaptive response has remained poorly characterized, and the underlying signaling pathways are unexplored. Here we show that hypertonicity induces submembranous de novo F-actin assembly concomitant with the peripheral translocation and colocalization of cortactin and the actin-related protein 2/3 (Arp2/3) complex, which are key components of the actin nucleation machinery. Additionally, hyperosmolarity promotes the association of cortactin with Arp2/3 as revealed by coimmunoprecipitation. Using various truncation or phosphorylation-incompetent mutants, we show that cortactin translocation requires the Arp2/3- or the F-actin binding domain, but the process is independent of the shrinkage-induced tyrosine phosphorylation of cortactin. Looking for an alternative signaling mechanism, we found that hypertonicity stimulates Rac and Cdc42. This appears to be a key event in the osmotically triggered cytoskeletal reorganization, because 1) constitutively active small GTPases translocate cortactin, 2) Rac and cortactin colocalize at the periphery of hypertonically challenged cells, and 3) dominant-negative Rac and Cdc42 inhibit the hypertonicity-provoked cortactin and Arp3 translocation. The Rho family-dependent cytoskeleton remodeling may be an important osmoprotective response that reinforces the cell cortex.

cell volume; cortactin translocation; Rac; Cdc42; actin-related protein 2/3


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ALTERATIONS IN CELL VOLUME occur under a variety of physiological and pathological conditions as a result of changes in the intra- or extracellular osmotic concentration. Such changes can be brought about by transport of metabolites and ions, polymerization or depolymerization of substrates, or exposure to anisoosmotic environments (14). Because extensive cell shrinkage or swelling represents an immediate threat to cellular integrity, several mechanisms have evolved to maintain homeostasis after volume perturbation. These adaptive responses either serve to restore near-normal cell volume or promote the reinforcement of the cell structure to withstand osmotic stress. Volume compensation is achieved through the activation of carriers and channels (37) and in the longer term by expression of genes that encode transporters and osmolyte-generating enzymes (50). The volume-dependent remodeling of the cytoskeleton is much less understood. Hyperosmotic stress has been documented to induce reorganization of the cortical actin network in yeast (7, 35) and Dictyostelium (1, 27, 40, 57), a process that has proven to be crucial for cell survival. In Dictyostelium, cell shrinkage caused redistribution of actin and actin-binding proteins including myosin and cofilin to the cortex, thereby providing increased physical resistance against osmotic shock. Besides this protective function, the cytoskeleton has been implicated as a volume sensor and mechanotransducer that may transmit signals to ion channels and transporters (16, 39). Further, the osmotically induced F-actin reorganization appears to be a central mechanism whereby hypertonicity inhibits neutrophil exocytosis and exerts clinically relevant anti-inflammatory effects (41).

Although osmotically induced changes in F-actin content or organization have been documented in different mammalian cells (for review, see Ref. 39), neither the mechanism of the volume-dependent F-actin response nor the underlying signaling events have been elucidated. The aims of the present work were to characterize shrinkage-induced molecular changes in the cortical skeleton and to explore the responsible signaling. We asked whether de novo actin nucleation contributes to osmotic remodeling of the cytoskeleton and whether recently discovered key components of the actin nucleation machinery, the actin-related protein 2/3 (Arp2/3) complex (17) and cortactin (54), could be involved in the volume-dependent cytoskeletal changes. This approach appeared attractive because our earlier studies have shown that cell shrinkage induces robust tyrosine phosphorylation of the F-actin binding protein cortactin (54, 55) through the activation of a novel volume-sensitive signaling cascade including Fyn and FER kinases (22, 24, 44). Furthermore, cortactin has recently emerged as an important organizer of cortical actin dynamics, and its accumulation is a strong indicator of de novo actin nucleation. Although its exact function is not well understood, cortactin has been implicated in cell motility (18), invasiveness (29, 38), shape determination (32), and vesicle-movement control (21). The biochemical basis of these diverse functions may be that cortactin has been found to bind and activate (in vitro) the Arp2/3 complex (48, 51), which is the major actin nucleating factor (17). In addition, cortactin appears to promote filament branching and inhibit debranching, thereby stabilizing the newly formed F-actin structure (51).

Cortactin harbors an NH2-terminal acidic tail (NTA) followed by five or six and a half tandem repeats (R), a proline-serine-threonine-rich region, a tyrosine-rich sequence, and a COOH-terminal SH3 domain (55). The NTA can bind to the Arp2/3 complex (48, 53), the R region binds and presumably cross links F-actin (19, 55), and tyrosine residues 421, 466, and 482 are the primary targets of Src family kinases and FER (18, 22), whereas the SH3 domain binds several membrane-associated proteins including dynamin (32) and the tight-junction protein ZO-1 (25). Cortactin is a target of both tyrosine kinases and the small GTPase Rac that has been shown to induce its translocation to the cell periphery (52). Interestingly, both of these pathways have been implicated in osmotic stress. However, the relationship between these signaling events and their exact role in the regulation of the functions of cortactin remain to be elucidated.

Here we examined the osmotic stress-induced reorganization of major cytoskeletal components: F-actin, cortactin, and the Arp2/3 complex. To define the underlying signaling, we tested the involvement of candidate mechanisms such as osmotically induced tyrosine phosphorylation or the hypertonic activation of Rac and Cdc42. We show that hyperosmotic stress induces cortical de novo actin assembly together with accumulation and association of cortactin and the Arp2/3 complex at the cell periphery. These events are independent of tyrosine phosphorylation. Importantly, we found that hypertonicity stimulates Rac and Cdc42 and induces translocation and colocalization of Rac with cortactin. Finally, we provide evidence that these small GTPases significantly contribute to the observed osmotic remodeling.


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Chemicals and antibodies. The enhanced chemiluminescence kit and the protein G-Sepharose beads were from Amersham Pharmacia Biotech. The protease inhibitor mixture containing 0.8 mg/ml benzamidine-HCl, 0.5 mg/ml aprotinin, 0.5 mg/ml leupeptin, 0.5 mg/ml pepstatin A, and 50 mM phenylmethylsulfonyl fluoride (PMSF) was obtained from PharMingen and was dissolved in pure ethanol. The Src family inhibitor 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2) was purchased from Calbiochem, Clostridium difficile toxin B (ToxB) was from TechLab, rhodamine-labeled and Alexa 488-labeled phalloidin were from Molecular Probes, and rhodamine-labeled G-actin was from Cytoskeleton. The following antibodies were used: monoclonal anti-phosphotyrosine (4G10), anti-cortactin (4F11), anti-Rac, and anti-Cdc42 were from Upstate Biotechnology; monoclonal (9E10) and polyclonal (rabbit) antibodies against c-Myc, polyclonal antibodies against Cdc42 and Arp3, and peroxidase-conjugated anti-goat IgM were from Santa Cruz Biotechnology; monoclonal anti-hemagglutinin (HA) antibody was from BabCo; FITC-labeled anti-goat and anti-rabbit IgGs and Cy3-labeled anti-mouse IgG were from Jackson Laboratories; and peroxidase-conjugated anti-mouse and anti-rabbit IgG were from Amersham Pharmacia Biotech.

Media. Bicarbonate-free RPMI 1640 was buffered with 25 mM HEPES to pH 7.4 (HPMI, osmolarity 290 ± 5 mosM). The isotonic sodium medium (Iso, 290 ± 5 mosM) consisted of (in mM) 130 NaCl, 3 KCl, 1 MgCl2, 1 CaCl2, 5 glucose, and 20 HEPES (pH 7.4). To obtain a hypertonic solution (Hyper), the Iso solution was supplemented with 300 mM sucrose to yield a final osmolarity of 600 ± 5 mosM.

Cell culture. In most experiments, we used Chinese hamster ovary (CHO) cells that stably express the rat sodium/proton exchanger 1 as in our previous studies (23). The behavior of these cells did not differ from wild-type CHO cells. The CHO cell line stably expressing wild-type, HA-tagged Cdc42 was a kind gift from Dr. R. A. Cerione (Cornell University; Ref. 10). In some experiments, the tubular epithelial cell line LLC-PK1 was used. CHO and LLC-PK1 cells were grown in alpha -minimal essential medium and in Dulbecco's modified Eagle medium, respectively. All media contained 25 mM NaHCO3 and were supplemented with 10% fetal calf serum and 1% antibiotic suspension (penicillin and streptomycin, Sigma). Cells were grown in a humidified air-CO2 (19:1 ratio) atmosphere at 37°C. CHO cells do not show any significant volume recovery after shrinkage under our experimental conditions.

Constructs and cell transfection. Green fluorescent protein (GFP)-cortactin constructs: the sequence coding for GFP was removed from pDXA-GFP plasmid and ligated into pcDNA 3.1(+) using existing HindIII/XbaI sites (Invitrogen). All cortactin constructs were amplified from pBluescript SK+ (Stratagene) using standard PCR procedures. Full-length cortactin was amplified using the primers sense, 5'-CAAAAGAGAAAGAATTCGAAAGCC and antisense, 5'-CTCTGGGTGGAATTCCTACTGCCG. The NH2 terminal cortactin construct (N-term, amino acids 1-334) was amplified using the primers sense, 5'-CAAAAGAGAAAGAATTCGAAAGCC and antisense, 5'-ATGGGGACAGAATTCTAATAGGC. The COOH terminal cortactin construct (C-term, amino acids 336-546) was amplified using the primers sense, 5'-TCTGCCTATCAGAATTCTGTC and antisense, 5'-CTCTGGGTGGAATTCCTACTGCCG. The cortactin SH3 domain construct (amino acids 458-546) was amplified using the primers sense, 5'-CAAGGCCTGACGAATTCATCA and antisense, 5'-CTCTGGGTGGAATTCCTACTGCCG. All primers have incorporated EcoRI sites for insertion into the EcoRI site of the pcDNA 3.1-GFP. Briefly, the modified vector was digested with EcoRI and gel purified, and the resulting fragment was treated with alkaline phosphatase (Promega) to prevent religation of the vector. Cortactin constructs were amplified by PCR, gel purified, digested with EcoRI, gel purified, and ligated into the pcDNA 3.1-GFP vector. Ligations were incorporated into DH5alpha , and the resulting colonies were screened for orientation of the insert by restriction digest. Clones with the insert in the proper orientation were confirmed by dideoxynucleotide sequencing. Construction of the other plasmids used in this study were described previously. The Myc-cortactin plasmid (19) encodes wild-type full-length murine cortactin tagged with the Myc epitope at its NH2 terminus. The Myc-cortactin Y421,466,482F plasmid encodes a mutant version of the full-length cortactin (designated as P-cortactin) in which the listed tyrosine residues were replaced by phenylalanines (18). The NTA encodes residues 1-80 of cortactin, whereas Delta NTA encodes a truncation mutant that lacks residues 1-68. These constructs as well as full-length cortactin were placed into a Tag5B vector (Invitrogen) that provided a COOH-terminal Myc tag (48). The constitutively active (Q61L) and dominant-negative (T17N) mutants of both Rac1 and Cdc42 are NH2 terminally Myc tagged (3, 56). Transient transfection with the corresponding plasmids was performed using FuGene reagent (Roche Molecular Biochemicals) according to the manufacturer's instructions. Routinely, cells were transfected with 1 µg of plasmid DNA per well (for 6-well plates) or 4-5 µg of DNA per 10-cm dish. The ratio of plasmid DNA to FuGene reagent was 1 µg to 2.5 µl, respectively. The details of cotransfection with two or three vectors are specified under the corresponding figures.

Preparation of cell extracts. Before the experiments, confluent cell cultures were kept in serum-free HPMI for 3 h. These quiescent cells were then preincubated in Iso medium for 10 min and subsequently subjected to various treatments as indicated. Unless otherwise stated, cells were treated with Iso or Hyper medium for 10 min. The medium was then aspirated and the cells were vigorously scraped into ice-cold lysis buffer (that contained 100 mM NaCl, 30 mM HEPES, 20 mM NaF, 1 mM EGTA, 1% Triton X-100, pH 7.5) supplemented with 1 mM Na3VO4, 1 mM PMSF, and 20 µl/ml protease inhibitor cocktail.

Immunoprecipitation and Western blotting. Lysates containing equal amounts of protein (0.8-1 mg) were clarified by centrifugation at 12,000× rpm for 10 min, precleared for 1 h using 35 µl of a 50% suspension of protein G-Sepharose beads, and then incubated with the corresponding antibodies for 1 h. Immuncomplexes were captured using 40 µl of a 50% suspension of protein G-Sepharose beads, and the beads were washed four times with lysis buffer that contained 1 mM Na3VO4. Immunoprecipitated proteins were diluted with Laemmli sample buffer, boiled for 5 min, and subjected to 8 or 10% SDS-PAGE as specified in the figures. The separated proteins were transferred to nitrocellulose using a Bio-Rad Mini Protean II apparatus. Blots were blocked in Tris-buffered saline that contained 5% bovine serum albumin (BSA) for 1 h and then incubated with the primary antibody for at least 1 h. Binding of the primary antibody was visualized by a 1:3,000 dilution of the relevant (anti-mouse, -rabbit, or -goat) peroxidase-coupled secondary antibody using the enhanced chemiluminescence method.

Rac and Cdc42 activity assays. The abundance of active (i.e., GTP-bound) small GTPase proteins was followed by pull-down assays (as described in Refs. 3, 28). Confluent cell cultures were serum deprived in HPMI medium for 3 h followed by a 10-min incubation in Iso medium. Subsequently, the medium was aspirated and replaced with either Iso or Hyper medium for the indicated times. Cells were then scraped in ice-cold magnesium lysis buffer [that contained 10% glycerol, 25 mM HEPES (pH 7.5), 150 mM NaCl, 1% Igepal CA-630, 10 mM MgCl2, 1 mM EDTA, and 1 mM Na3VO4] supplemented with 1 mM PMSF and 20 µl/ml protease inhibitor cocktail, and the lysates were precleared by brief centrifugation (1 min at 12,500× rpm). To capture active Rac and Cdc42, the supernatants were immediately mixed with 10 µl of a 50% suspension of glutathione beads covered with a fusion protein (approx 10 µg) composed of glutathione S-transferase (GST) and the p21-binding domain of the p21-activated kinase (PAK, Upstate Biotechnology). After a 30-min rotation at 4°C, the beads were washed four times with magnesium lysis buffer, suspended in Laemmli sample buffer, and boiled for 5 min. Precipitated proteins were subjected to electrophoresis on 15% SDS-polyacrylamide gels, which was followed by Western blotting using anti-Rac or anti-Cdc42 antibodies. To obtain controls for active and inactive GTPases, lysates from untreated cells were supplemented with 0.1 mM GTPgamma S or 1 mM GDP, respectively, and incubated for 15 min. Nucleotide binding was locked by adding 60 mM MgCl2 to the lysates, and the samples were analyzed.

Immunofluorescence microscopy. Confluent cultures grown on 25-mm coverslips were serum deprived for 3 h in HPMI, preincubated for 10 min in Iso medium, and treated as specified in the figure captions. Cells were fixed for 30 min in Iso or Hyper medium (as used for the experiment) supplemented with 4% paraformaldehyde. The coverslips were extensively washed with PBS, incubated with 100 mM glycine in PBS for 10 min, permeabilized with 0.1% Triton X-100 in PBS for 20 min, and then blocked with 3% BSA or 1% donkey serum in PBS for 1 h. Samples to be stained with the anti-Arp3 antibody were fixed with ice-cold methanol for 5 min and blocked as described. Subsequently, the samples were incubated with primary antibodies for 1 h, washed with PBS, and incubated with fluorescently labeled secondary antibodies for 1 h. For the detection of F-actin, fixed and permeabilized cells were incubated with rhodamine-phalloidin or Alexa 488-phalloidin. The coverslips were washed and mounted on glass slides using mounting solution (DAKO). The staining was visualized using either a Leica DM1RB fluorescence microscope (×100 objective) coupled to a Micromax cooled charge-couple device (CCD) camera (Princeton Instruments) driven by WinView software or a Nikon Eclipse TE200 microscope (×100 objective) coupled to a Hamamatsu cooled CCD camera (C4742-95) controlled by Simple PCI software. Where indicated, confocal images were obtained using a Zeiss LSM 510 confocal microscope (×100 objective) and LSM 510 software. Size bars corresponding to 10 µm were added to images where applicable.

Incorporation of rhodamine-G-actin. Confluent cultures grown on 25-mm coverslips were serum deprived for 3 h in HPMI, preincubated for 10 min in Iso medium, and treated either isotonically or hypertonically for 1.5 min or with phorbol 12-myristate 13-acetate (PMA) for 3 min. After treatment, cells were incubated for 1 min at 37°C in permeabilization buffer that contained 138 mM KCl, 10 mM PIPES, 3 mM EGTA, 4 mM MgCl2, 1% BSA, 0.025% saponin, 0.1 mM ATP, and 0.5 µM rhodamine-G-actin (5, 6). The permeabilization buffer was then aspirated, and the cells were briefly washed and immediately fixed with 4% paraformaldehyde in PBS. After fixation, the cells were washed extensively with PBS and in some cases were then stained for F-actin using Alexa 488-phalloidin as described (see Immunofluorescence microscopy). Incorporated rhodamine-G-actin and F-actin were viewed by fluorescence microscopy. To control for surface binding of rhodamine-G-actin, saponin was left out of the permeabilization buffer.

Protein assays. Protein concentrations were determined by bicinchoninic acid assay (BCA Assay, Pierce) using BSA as a standard.

Densitometry. Quantification of the bands was performed using a Bio-Rad GS-690 Imaging Densitometer and the Molecular Analyst program as described previously (24).

Data are presented as representative immunoblots or photomicrographs of at least three similar experiments or as the means ± SE of the number of experiments indicated (n).


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Hypertonicity induces actin skeleton remodeling and de novo F-actin assembly at the cell periphery. To assess the effect of hyperosmotic stress on the actin skeleton, CHO cells were exposed to iso- or hyperosmotic conditions, and after fixation the F-actin structure was visualized using rhodamine-phalloidin staining. A short exposure to moderate hypertonicity (10 min, extra 300 mosM) induced major structural changes in F-actin organization that manifested in a reduction of the number and width of stress fibers concomitant with a substantial increase in peripheral F-actin staining (Fig. 1A). To address the mechanism underlying the observed peripheral F-actin accumulation, we wished to establish whether hypertonicity induced an increase in actin nucleation at the periphery. To this end, we used a technique that allows spatial resolution of active nucleation sites, which was successfully applied to detect epidermal growth factor-induced peripheral F-actin assembly (5, 6). The basis of this method is that in cells briefly permeabilized with a mild detergent, active nucleation sites can be visualized due to an enhanced ability to incorporate exogenously supplied labeled monomeric actin (G-actin). We therefore exposed CHO cells for a brief (1.5-min) iso- or hypertonic treatment, which was followed by permeabilization with 0.025% saponin in the presence of rhodamine-G-actin. After a 1-min incubation, the cells were washed and fixed. First, we had to test whether the method exclusively detects intracellular staining. Figure 1B (top) shows that absolutely no staining was present in nonpermeabilized cells, which clearly indicates that labeling cannot be attributed to the surface attachment of G-actin. In isotonically treated and then permeabilized cells, diffuse, finely punctate fluorescence was observed that was most pronounced in the central areas (Fig. 1B, Iso). In contrast, most hyperosmotically treated and then permeabilized cells showed distinct peripheral labeling in the form of sharp lines at the cell boundary (see arrows on Fig. 1B, Hyper). As a positive control, we used PMA to induce ruffling, because this process is characterized by de novo actin assembly. In agreement with this expectation, PMA caused increased peripheral G-actin incorporation (Fig. 1B, PMA) that was best visible at the free edges of the cells. To examine whether the added G-actin was indeed incorporated into F-actin, we performed similar experiments, but at this time the cells were also stained with Alexa 488-phalloidin. Consistent with our findings above, peripheral rhodamine-G-actin staining was very weak under isotonic conditions, although phalloidin clearly visualized some F-actin at the periphery as well as abundant stress fibers (Fig. 1C, Iso). In contrast, areas that showed intense peripheral rhodamine labeling in hypertonically treated cells precisely colocalized with strong peripheral Alexa 488-phalloidin (F-actin) staining. This finding implies that the rhodamine-positive lines at the periphery correspond to the incorporation of G-actin into newly generated F-actin (Fig. 1C, Hyper, see arrows). Taken together, these experiments suggest that hyperosmolarity promotes de novo F-actin assembly at the periphery.


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Fig. 1.   Hyperosmolarity induces actin skeleton reorganization and facilitates de novo F-actin assembly at the cell periphery. A: Chinese hamster ovary (CHO) cells grown on glass coverslips were serum deprived for 3 h in bicarbonate-free RPMI 1640 buffered with 25 mM HEPES to pH 7.4 and preincubated with an isotonic solution (Iso) for 10 min. Subsequently the medium was exchanged for either the same Iso solution or a hypertonic solution (Hyper). To achieve hypertonicity, the Iso solution was supplemented with 300 mM sucrose. After a 10-min treatment, cells were fixed and stained for F-actin using rhodamine-labeled phalloidin. Note that hypertonicity induces peripheral accumulation of F-actin and reduces stress fibers. B: serum-deprived CHO cells were treated with either Iso or Hyper solution for 1.5 min or with 100 nM phorbol 12-myristate 13-acetate (PMA) for 3 min. Cells were permeablized in the presence of rhodamin-labeled G-actin for 1 min, nonincorporated rhodamine-G-actin was briefly washed away, and the samples were immediately fixed. As a control, CHO cells treated with Iso solution were incubated with rhodamine-G-actin in the absence of the detergent saponin (No saponin, top) and viewed with fluorescent light (left) and phase contrast (right). Incorporated G-actin in permeablized cells (bottom) was visualized by fluorescent microscopy. Note the increase in G-actin incorporation at the periphery in hypertonically treated cells and at the ruffles in PMA-stimulated cells (see arrows). C: CHO cells were treated isotonically or hypertonically for 1.5 min, permeablized in the presence of rhodamine-G-actin, briefly washed, and fixed and stained for F-actin using Alexa 488-labeled phalloidin. Incorporated G-actin (top) and F-actin (bottom) were visualized by fluorescent microscopy. Notice that peripherally incorporated G-actin colocalizes with F-actin staining in hypertonically treated cells (see arrows). Size bars, 10 µm.

Hypertonicity induces cortactin translocation to cell periphery independent of tyrosine phosphorylation. Recent studies have implicated cortactin as an indicator and positive regulator of de novo actin nucleation (17). Moreover, our previous work has shown that it is a volume-sensitive protein because it undergoes robust tyrosine phosphorylation upon cell shrinkage (22, 24). To assess whether cortactin is involved in osmotically induced cortical cytoskeleton remodeling, we investigated whether a decrease in cell volume affects its distribution. Under isoosmotic conditions, cortactin showed a finely punctate, even distribution throughout the cytosol with clear nuclear exclusion (Fig. 2A, Iso). Hyperosmotic treatment caused a marked change in cortactin localization: the staining in the cell interior became reduced while increased labeling occurred at the cell periphery, which suggests that cortactin moved to the submembranous area (Fig. 2A, Hyper). Osmotic stress induced similar cortactin reorganization both in fibroblast-type CHO cells and epithelial LLC-PK1 cells (Fig. 2A). In hypertonically treated cells, the fine granularity of the cytosol disappeared, and occasionally bigger aggregates were also observed. Cortactin peripheralization was rapid and sustained; it was usually detectable within 2-3 min after hyperosmotic exposure and persisted throughout the duration of the osmotic stress. To test whether the observed cortactin redistribution was not an artifact due to altered epitope accessibility in the shrunken cells, we transfected CHO cells with a construct encoding for full-length GFP-cortactin and directly followed the distribution of the expressed protein. Under isotonic conditions, GFP-cortactin showed perinuclear/cytosolic localization without any enrichment at the periphery, whereas hyperosmotic treatment caused marked GFP-cortactin accumulation along the plasma membrane (Fig. 2B). Because cell shrinkage changes cell geometry, it was important to test whether the observed redistribution was a real phenomenon and not simply an optical artifact caused by the altered cell shape. To assess this, we transfected cells with a construct encoding five GFP molecules coupled together (5GFP). We chose this approach because 5GFP, in contrast to the smaller GFP, showed similar nuclear exclusion as endogenous or GFP-labeled cortactin. After iso- or hypertonic treatment, we stained the cells for endogenous cortactin. Figure 2C shows that osmotic stress failed to affect the distribution of 5GFP (Fig. 2C, top), whereas it caused peripheral accumulation of endogenous cortactin in the same cell. Thus the observed cortactin redistribution reflects real translocation and cannot be attributed to an optical artifact due to cell shrinkage.


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Fig. 2.   Hypertonicity induces cortactin translocation to the cell periphery. A: CHO cells or LLC-PK1 cells were treated isotonically or hypertonically. After a 10-min treatment, cells were fixed and immunostained for cortactin. Note that under isotonic conditions, cell boundaries are hardly visible because cortactin is evenly distributed excluding only the nuclear region. Hypertonic treatment induces a marked increase in the peripheral staining and permits clear distinction of the cell borders. B: CHO cells were transiently transfected with green fluorescent protein (GFP)-tagged, full-length cortactin (1 µg of DNA/well). After 48 h, cells were serum deprived, treated with Iso or Hyper solutions for 10 min, and fixed. C: CHO cells were transfected with a construct of five GFP molecules coupled together (5GFP), treated iso- or hypertonically for 10 min, fixed, and stained for endogeneous cortactin.

Next we examined whether tyrosine phosphorylation of cortactin was required for its redistribution using both genetic and pharmacological approaches. First, we transfected CHO cells either with wild-type Myc-tagged cortactin or with a Myc-tagged cortactin mutant (P-cortactin) in which three critical tyrosine residues (421, 466, and 482) were replaced with phenylalanines (18). Consistent with our earlier findings (22), osmotic stress induced tyrosine phosphorylation in the wild-type cortactin but failed to do so in P-cortactin (Fig. 3A). To compare the localization of the wild-type and P-cortactin, cells were transfected with the corresponding constructs and 48 h later were treated iso- or hypertonically and then fixed and stained with an anti-Myc antibody. Under isotonic conditions, the distributions of Myc-cortactin and Myc-P-cortactin were similar and comparable to that of the endogenous protein (Fig. 3B, a and c). Moreover, upon hyperosmotic treatment, both the wild-type and the mutant cortactin efficiently translocated to the cell periphery (Fig. 3B, b and d). Because cortactin has been suggested to form an oligomer (48), it was conceivable that the endogenous protein translocates in a tyrosine phosphorylation-dependent manner and subsequently could recruit nonphosphorylated cortactin molecules. To address this possibility, we tested whether the Src family inhibitor PP2 that completely abolishes osmotic cortactin phosphorylation (Fig. 3A and Ref. 24) alters the translocation of the endogenous protein. Figure 3C shows that PP2 failed to inhibit cortactin movement. Consistent with this, we found that PP2 did not affect the osmotically induced F-actin accumulation either (not shown). Together these findings indicate that osmotic cortactin phosphorylation is not required for the concomitant cortactin translocation and point to the involvement of other signaling pathways and/or the shrinkage-induced mechanical reorganization in the cytoskeleton.


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Fig. 3.   Hyperosmolarity-induced cortactin translocation is independent of cortactin tyrosine phosphorylation. A: CHO cells grown on 10-cm dishes were transiently transfected (4 µg of DNA/dish) with constructs encoding either Myc-cortactin or the mutant Myc-P-cortactin in which the major target tyrosine residues of the Src family were replaced by phenylalanines (Y421, 466, 482F). Cells were treated 48 h later with Iso (I) or Hyper (H) solutions for 10 min. Where indicated, cells were preincubated with 10 µM 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2) for 10 min followed by iso- or hypertonic treatment in the presence of the drug. Subsequently the cells were lysed and processed for immunoprecipitation with an anti-Myc antibody. Precipitates were probed with anti-phosphotyrosine antibody (anti-PY) and reprobed with anti-Myc antibody (anti-Myc). B: CHO cells grown on glass coverslips were transiently transfected with 1 µg of either Myc-cortactin (a and b) or Myc-P-cortactin (c and d). Cells were treated 48 h later with Iso (a and c) or Hyper (b and d) solutions and fixed and stained for Myc. C: CHO cells (e and f) and LLC-PK1 cells (g and h) were preincubated with 10 µM PP2 for 10 min and exposed to Iso or Hyper solutions containing the same PP2 concentration. After treatment, cells were fixed and stained for endogenous cortactin. WB, Western blot.

NH2-terminal half of cortactin is sufficient for hypertonicity-induced translocation. To gain insight into the structural requirements of cortactin redistribution and its relationship to actin remodeling, we explored which domains of cortactin were necessary for shrinkage-induced translocation. Both NH2- and COOH-terminal cortactin domains have been reported to bind to membrane-associated proteins: the NTA and R regions can bind to the Arp2/3 complex and F-actin, respectively (48, 53, 55), whereas the COOH-terminal SH3 domain can associate with dynamin (32). To visualize movements of cortactin and its various domains independent of the presence of a given epitope, we transfected CHO cells with constructs encoding for GFP-tagged, full-length cortactin or various truncation mutants. Subsequently, cells were treated iso- or hypertonically and stained with rhodamine-phalloidin, and the distribution of GFP-cortactin proteins and F-actin were visualized using dual-wavelength confocal microscopy. As expected, hypertonicity induced the accumulation of full-length cortactin at the periphery (Fig. 4A, Full-length, GFP) where it colocalized with the thick submembranous F-actin ring (see the yellow color in Fig. 4A, Merge). The NH2-terminal half (N-term) harboring the NTA and R domains behaved similarly in that it showed even cytosolic distribution under isotonic conditions, whereas hypertonicity induced its translocation and colocalization with F-actin (Fig. 4A, N-term). In contrast, the COOH-terminal half (C-term) and the isolated SH3 domain failed to translocate upon osmotic stress. GFP alone also remained unaffected. These findings indicate that the NH2-terminal half is both necessary and sufficient for osmotic translocation of cortactin, whereas the COOH terminus does not appear to participate in this phenomenon.


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Fig. 4.   The NH2-terminal half of cortactin is sufficient for the osmotically induced translocation. A: CHO cells were transfected with full-length GFP-cortactin, GFP alone, or the following GFP-tagged cortactin truncation mutants: the NTA and R region (N-term); the helix, Pro-, Ser-, and Thr-rich, and SH3 domains (C-term); or the SH3 domain alone (SH3). Two days later, cells were treated iso- or hypertonically and fixed and stained for F-actin using rhodamine-phalloidin. Green (GFP-constructs) and red (F-actin) fluorescence was visualized using dual-wavelength laser-scanning confocal microscopy. In the merged image, the yellow regions indicate colocalization of the green and red signals. Note that only full-length and N-term cortactin are able to translocate to the periphery (see arrows) and colocalize with F-actin. B: CHO cells were transfected with the following COOH terminally Myc-tagged constructs: full-length cortactin (top), NTA region of cortactin (middle), or a cortactin trunctation mutant in which the NTA region was deleted (Delta NTA, bottom). After transfection, cells were challenged iso- or hypertonically and fixed and stained for Myc.

Because the NH2 terminus is composed of two functionally different domains (NTA and R), we investigated the potential role of each of these. To this end, we used two Myc-tagged constructs: one encoding almost the full NTA (amino acids 1-80) and the other lacking the first 68 amino acids of the NH2 terminus (Delta NTA). As illustrated in Fig. 4B, both truncation mutants showed redistribution to the cell borders in hypertonically treated cells. However, the translocation appeared less pronounced inasmuch as it was more diffuse under the membrane and usually restricted to smaller areas. These experiments suggest that either the isolated NTA or the R region is capable of translocating to the cell periphery where these constructs presumably interact with proteins of the actin nucleation machinery. Nevertheless, the translocation or retention seems to be more efficient when both domains are present.

Hyperosmolarity induces peripheral accumulation of Arp3 and its association with cortactin. The increased G-actin incorporation together with the observed peripheral cortactin accumulation strongly suggests that osmotic stress enhances de novo F-actin nucleation activity under the membrane. Because the Arp2/3 complex is a prime regulator of this process and can directly bind to the NTA domain, we examined whether it is also recruited upon hyperosmotic stimulation. In CHO cells, Arp3 showed an even cytosolic distribution under isotonic conditions. Hypertonicity induced a marked increase in the labeling at the periphery that was most prominent along the cell-cell contacts (Fig. 5A, top, see arrows). In addition, the osmotically redistributed Arp3 appeared to colocalize with cortactin as verified by double immunostaining (Fig. 5A, bottom). The phenomenon was similar in LLC-PK1 cells. In these cells, cortical areas showed weak Arp3 accumulation even under isotonic conditions, which was dramatically increased upon hyperosmotic exposure (Fig. 5B). Next we investigated whether an association between Arp2/3 and cortactin could be verified by biochemical means. We immunoprecipitated Arp3 from lysates of iso- or hypertonically treated cells and probed the precipitates with an anti-cortactin antibody. After isotonic treatment, only a marginal amount of cortactin was present in the Arp3 precipitates. In contrast, after hyperosmotic exposure, the Arp3 antibody pulled down a substantial amount of cortactin, which indicates that hypertonicity promoted the association of these proteins (Fig. 5C). The 80- and 85-kDa isoforms of cortactin were equally increased in the precipitate. To test the effect of tyrosine phosphorylation on the relationship between Arp3 and cortactin, similar experiments were performed using PP2-treated cells. The inhibitor increased the coprecipitation of Arp3 and cortactin in the isotonic samples and potentiated the association after hypertonic exposure. These findings show that the tyrosine phosphorylation of cortactin is not required for its association with Arp3. Rather, tyrosine phosphorylation might be a negative regulator that promotes the dissociation of the complex. Collectively, these results show that hypertonicity induces major changes in the cortical skeleton and causes peripheral accumulation and complex formation between F-actin, cortactin, and Arp2/3.


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Fig. 5.   Hypertonicity induces the translocation of actin-related protein 3 (Arp3) and its association with cortactin. CHO (A) or LLC-PK1 (B) cells were treated iso- or hypertonically, fixed with methanol, and stained for Arp3 and cortactin (A) or Arp3 alone (B). CHO cells (C) were preincubated in the absence or presence of PP2 and treated with iso- or hypertonic solutions. Cells were lysed, and Arp3 was immunoprecipitated (IP) from the clarified lysates. Precipitates were processed for Western blotting and probed with the anti-cortactin antibody. To assess whether the initial Arp3 and cortactin contents were the same, samples from the Triton X-soluble fraction were run on 8 and 10% SDS-PAGE gels and were Western blotted with anti-cortactin and anti-Arp3 antibodies, respectively. For Iso and hyper treatments, duplicates are shown.

Osmotic stress activates small GTPases Rac and Cdc42. The Rho family small GTPases, Rac and Cdc42, play an essential role in the organization of the cortical cytoskeleton (34), and we have recently shown that hypertonicity increased the activity of these small GTPases in suspended neutrophils (28). Because activated Rac has been shown to induce cortactin translocation to the cell periphery in fibroblasts (52), it was conceivable that Rac and/or Cdc42 might mediate the osmotic translocation of cortactin and Arp2/3. To address this hypothesis, we initially examined whether hyperosmotic stress induces the activation of these small GTPases in CHO cells. We applied two approaches. First, we tested the effect of hypertonicity on the intracellular localization of Rac and Cdc42, because membrane translocation of these small G proteins is thought to be a strong though indirect indicator of activation (45). The cells were exposed to Iso or Hyper solutions for 10 min, and after fixation the small GTPases were visualized by immunostaining. In serum-starved cells under isotonic conditions, Rac was evenly distributed in the cytosol in a finely punctate manner. Hypertonicity caused a marked increase in peripheral labeling that was manifested in many cells as enhanced Rac staining in a narrow line along the cell border (Fig. 6A, top).


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Fig. 6.   Rac and Cdc42 are activated by hyperosmolarity. A: wild-type CHO cells (top) or CHO cells stably expressing hemagglutinin (HA)-tagged wild-type Cdc42 (bottom) were exposed to Iso or Hyper solutions and fixed and stained for Rac (top) or Cdc42 (bottom) using anti-Rac and anti-HA antibodies, respectively. Notice that hyperosmolarity caused redistribution of Rac and Cdc42 to the cell periphery. B: CHO cells were treated isotonically (Iso) or hypertonically (Hyp) for 10 min or with 100 nM PMA for 5 min. Additionally, CHO cells were treated with Hyper solution for 10 min and were then returned to isotonicity for 1 min (Hypright-arrowIso 1') or 5 min (Hypright-arrowIso 5'). Cells were lysed and the active forms of Rac and Cdc42 were pulled down from the lysates using glutathione S-transferase (GST)-p21-binding domain (PBD) beads. To check for the initial equality of small GTPases in the samples, aliquots of the cell lysates were obtained before addition of the GST-PBD beads. Precipitates (Active Rac/Cdc42) and cell lysates (Total Rac/Cdc42) were subjected to SDS-PAGE (15% gel) and Western blotting using anti-Rac and anti-Cdc42 antibodies. To confirm that the GST-PBD beads specifically precipitated the active forms of Rac and Cdc42, positive and negative controls were performed on cell lysates using GTPgamma S and GDP, respectively, and were analyzed as above. C: graph represents the hypertonicity-induced relative increase in GTP-bound Rac and Cdc42 as determined by densitometric analysis of 6 and 10 separate experiments, respectively. D: CHO cells were treated with Iso solution for 10 min or with Hyper solution for the indicated times. Activities of Rac and Cdc42 were determined as described in B.

To visualize Cdc42, we used a CHO cell line that stably expresses the HA-tagged, wild-type Cdc42, because the available anti-Cdc42 antibody was not optimal for immunofluorescence studies (Fig. 6A, bottom). Under resting conditions, Cdc42 was evenly scattered in the cytosol in fine punctate aggregates, and it was also present occasionally in ruffles. Upon hypertonic treatment, the labeling accumulated in a narrow region along the periphery in most cells. This morphological finding is consistent with a recent biochemical approach which shows that osmotic shock-induced enrichment of Cdc42 in a particulate fraction (42).

Next we used a recently developed pull-down assay to test whether the observed translocation was indeed coincident with the activation of these small GTPases. Lysates obtained from iso- and hypertonically treated cells were incubated with a fusion protein composed of GST and the p21-binding domain (PBD) of PAK, which is a downstream effector of both Cdc42 and Rac (3). PBD binds the active (GTP-bound) form of the small GTPases but fails to capture the inactive (GDP-bound) form (Fig. 6B, lanes GTPgamma S and GDP). The fusion protein was adsorbed to GSH beads, and the bead-associated proteins were subjected to SDS-electrophoresis followed by Western blotting using anti-Cdc42 and anti-Rac antibodies. Hypertonic treatment (+300 mosM for 10 min) caused 3.33 ± 1.04-fold (n = 6) and 2.46 ± 0.32-fold (n = 10) increases in the activity of Rac and Cdc42, respectively (Fig. 6C). The magnitude of the hypertonic effect was comparable to that of PMA, which is a known activator of Rac (3). The osmotic stimulation of the small GTPases was rapid and sustained: an increase in activity was detectable after 1 min, continued to rise over ~10 min, and remained elevated throughout the hypertonic exposure (Fig. 6D). Upon restoration of isotonicity, GTP-Rac and GTP-Cdc42 decreased to prestimulus levels within a few minutes (Fig. 6B, Hypright-arrowIso). Taken together, these observations indicate that hyperosmotic stress induces the translocation and concomitant activation of Rac and Cdc42.

Involvement of Rac and Cdc42 in hypertonicity-triggered cortactin and Arp3 translocation. The next set of experiments was carried out to test whether active Rac and Cdc42 could indeed induce cortactin translocation in CHO cells and to establish whether the active small GTPases colocalized with cortactin at the periphery. The cells were transiently cotransfected with wild-type GFP-tagged cortactin and Myc-tagged constitutively active (CA, Q61L) Rac or Cdc42. After 48 h, the cells were stained for the Myc epitope and viewed by confocal microscopy (Fig. 7A). Alternatively, cells were cotransfected with CA Rac or CA Cdc42 along with GFP (to identify the successfully transfected cells) and were stained for endogenous cortactin (not shown). Active Rac caused the formation of prominent ruffles and lamellipodia in 91% of the cells (n = 187 cells). These structures showed strong staining for active Rac that colocalized with GFP-cortactin (Fig. 7A, top). The latter finding was intriguing, because in previous studies the active Rac-induced translocation of cortactin was not shown to be accompanied by the translocation of Rac per se (see DISCUSSION). CA Cdc42 had similar but less-prominent effects. In our cells, the long-term expression of Cdc42 did not result in extensive filopodia formation; rather, it induced the formation of rufflelike structures at the periphery as reported in other fibroblasts as well (26). Interestingly, CA Cdc42 was also present in the ruffles (Fig. 7A, bottom). These morphological features were observed in 56% of the CA Cdc42-expressing cells (n = 176), i.e., these changes occurred less frequently than in CA Rac-transfected cells. However, in all cases when CA Cdc42 showed enrichment at these peripheral structures, cortactin colocalized with it (Fig. 7A, bottom). Given the similarity between the morphological changes induced by the two small GTPases, it is conceivable that under our conditions Cdc42 exerted its effect predominantly through the activation of Rac (26, 34). Together these results show that CA Rac and CA Cdc42 are able to induce cortactin translocation to the periphery in CHO cells, and the small GTPases themselves are present in the same structures as cortactin. Rac, however, appears to be more efficient.


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Fig. 7.   Effect of constitutively active (CA) Rac and Cdc42 and hypertonicity on the distribution of cortactin and the small GTPases. A: CA forms of Rac and Cdc42 translocate cortactin and colocalize with cortactin at the cell periphery. CHO cells were transfected with 1 µg of GFP-cortactin along with either 2 µg of Myc-tagged Q61L-Rac (CA, top) or 2 µg of Myc-tagged Q61L-Cdc42 (CA, bottom). After 48 h, cells were fixed, stained for Myc, and viewed with dual-wavelength confocal microscopy. Myc-staining (red, left), GFP-cortactin (green, middle), and merged images (right). Yellow areas represent colocalization. Arrows indicate membrane ruffles in which GFP-cortactin and CA Rac or Cdc42 colocalize. B: hyperosmolarity induces colocalization of Rac and GFP-cortactin at the periphery. CHO cells were transfected with GFP-cortactin, challenged isotonically (top) or hypertonically (middle and bottom), immunostained for endogenous Rac, and viewed as in A. Rac (red, left), GFP-cortactin (green, middle), merged images (right). Arrows indicate areas where Rac and GFP-cortactin colocalize. Typically the osmotic stress-induced translocation of cortactin and Rac occurred along the entire periphery (middle). Less frequently, translocation and colocalization of these proteins were observed in distinct rufflelike structures (bottom).

We then performed experiments to directly assess the role of these small GTPases in osmotic cortactin redistribution. First, we tested whether Rac and cortactin are attracted to the same location upon hyperosmotic exposure. Cells were transfected with GFP-cortactin and, after iso- or hyperosmotic treatment, were stained for endogenous Rac before being examined by confocal immunofluorescence microscopy. Although the overall pattern of Rac and cortactin distribution was similar in isotonically treated cells, no colocalization occurred between these molecules as verified by the merged confocal images (Fig. 7B, Iso). This finding suggests that these proteins are not present in the same macromolecular complex. In contrast, after hypertonic exposure, we observed a high level of colocalization of Rac and cortactin at the cell periphery (Fig. 7B, Hyper). In many cells, increased Rac and GFP-cortactin labeling occurred along the entire boundary (Fig. 7B, middle), whereas in a smaller number of cells, augmented accumulation of these proteins was restricted to peripheral rufflelike structures (Fig. 7B, bottom). Moreover, peripheral Rac accumulation appeared to be enhanced in cortactin-overexpressing cells, which raises the possibility that Rac-induced cortactin translocation might facilitate the translocation of Rac itself via a positive-feedback mechanism.

To substantiate the role of osmotic activation of small GTPases in cortactin translocation, we coexpressed the Myc-tagged, dominant-negative (DN) versions of Rac and Cdc42 and followed the effect of hypertonicity on cortactin translocation in these cells. The DN constructs had no effect on the basal cortactin distribution but did inhibit the hypertonicity-induced cortactin translocation. Peripheral cortactin localization was observed in 92% of hypertonically treated control cells (n = 200), whereas it was seen only in 42.7% of the transfected cells (n = 138) providing ~50% inhibition (Fig. 8A). The inhibitory effect was even more pronounced in LLC-PK1 cells, where 70.0% of the DN Rac/Cdc42-expressing cells (n = 130) did not show cortactin translocation (Fig. 8B). Similar results were obtained when only DN Rac was transfected (not shown). Importantly, these effects were not due to a loss of cell viability nor a manifestation of general refractoriness to cell shrinkage, because the DN Rac-transfected cells retained the cytoplasmic pH indicator 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein and responded to a hypertonic challenge with activation of the Na+/H+ exchanger (K. Szászi, unpublished observation). Together these data suggest that Rac plays an important but not exclusive role in the osmotically induced translocation of cortactin.


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Fig. 8.   Dominant negative (DN) forms of Rac and Cdc42 inhibit the hyperosmotic stress-induced cortactin and Arp3 translocation. CHO cells or LLC-PK1 cells were cotransfected with 1 µg of Myc-tagged T17N-Rac and 1 µg of Myc-tagged T17N-Cdc42. After 48 h, cells were challenged with Iso or Hyper solutions and fixed and stained for either Myc and cortactin or Myc and Arp3. Identical symbols represent identical cells on the corresponding Myc and cortactin or Arp3 images. A and B: cytosolic cortactin labeling is preserved under isotonic conditions in both nontransfected and DN Rac/Cdc42-expressing cells. Hyperosmotically induced cortactin translocation was obvious in >90% of the nontransfected control cells. In contrast, in DN Rac/Cdc42-expressing cells, peripheral accumulation of cortactin is absent or reduced after hypertonic stimulation. Complete inhibitory effect (preserved cytosolic staining without peripherialization) was observed in 57.3% (n = 138) and 70.0% (n = 130) in CHO and LLC-PK1 cells, respectively. C: DN Rac and Cdc42 prevented the peripheral accumulation of Arp3 in 83% (n = 100) of hypertonically treated cells. To avoid any ambiguity that might arise from assessing peripheral accumulation at the border between a DN small GTPase-expressing and nonexpressing cell, we preferentially counted those cells that had transfected neighbors.

To assess whether the hyperosmotically induced Arp3 recruitment also depended on small GTPase activation, we expressed DN Rac/Cdc42 in LLC-PK1 cells and analyzed Arp3 distribution under iso- and hypertonic conditions. Although the DN constructs had no significant impact on Arp3 distribution under isotonic conditions, the constructs exerted a dramatic inhibitory effect on the hypertonicity-provoked Arp3 translocation. Arp3 translocated in virtually all control cells but failed to accumulate at the periphery in 83% of DN Rac/Cdc42-expressing cells (n = 100; Fig. 8C).

To address whether the DN constructs reduced cortactin translocation by interfering with the osmotically induced F-actin reorganization, we transfected cells with GFP-cortactin alone (Fig. 9A) or together with DN Rac/Cdc42 (Fig. 9B) and visualized F-actin with rhodamine-phalloidin. As shown in Fig. 9B, under isotonic conditions, many DN Rac/Cdc42-transfected cells showed a disorganized F-actin staining and normal cortactin distribution. Upon hypertonic challenge, many DN Rac/Cdc42 expressors failed to develop the typical peripheral F-actin deposition, and in these cells cortactin remained in the cell interior (Fig. 9B). However, the cells showed heterogeneity, and we observed transfected cells in which hypertonicity still induced actin and cortactin movement to the periphery. Thus we infer from these experiments that DN Cdc42 and DN Rac interfere with osmotic reorganization of F-actin and cortactin relocalization. On the other hand, the inhibition is not complete, which suggests the involvement of additional mechanisms.


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Fig. 9.   Parallel determination of the effect of DN Rac and Cdc42 on the organization and osmotic remodeling of F-actin and on cortactin redistribution. CHO cells were transfected with 1 µg of GFP-cortactin alone (A) or together with 1 µg of DN Rac plus 1 µg of DN Cdc42 (B). After iso- or hypertonic treatment, cells were fixed and stained for F-actin using rhodamine-phalloidin. F-actin (A and B, top) and GFP-cortactin (A and B, bottom) were then visualized using the corresponding excitation filters. Identical symbols represent identical cells on corresponding cortactin and actin images; arrow shows the translocation of GFP-cortactin in a control cell.

Role of small GTPases in osmotic phosphorylation of cortactin. Our data show that translocation of cortactin does not require tyrosine phosphorylation. However, it is conceivable that phosphorylation (at least in part) occurs as a result of translocation to the periphery where cortactin might be phosphorylated by membrane-associated tyrosine kinases. In favor of this possibility, Src was shown to translocate to the periphery in a Rac-dependent manner in platelet-derived growth factor-stimulated cells (12). To address whether the activation of the small GTPases played a role in the osmotic phosphorylation of cortactin, we treated the cells with ToxB, a potent inhibitor of the Rho family small GTPases (20). A 4-h exposure resulted in cell rounding in 100% of the cells, which clearly implies that the drug effectively penetrated the cells and exerted its effect on the Rho family. Control and ToxB-treated cells were then challenged with Iso or Hyper solutions, and cortactin immunoprecipitates obtained from these samples were analyzed by Western blotting using anti-phosphotyrosine antibody. Densitometric analyses of three similar experiments showed that ToxB caused a slight but consistent reduction (average of 30%) in the hypertonic tyrosine phosphorylation (Fig. 10A), which suggests that the activation of small GTPases might facilitate cortactin phosphorylation. Nevertheless, in each of the three experiments, strong cortactin phosphorylation occurred in the presence of ToxB. To substantiate these pharmacological data, cells were cotransfected with constructs encoding for Myc-tagged DN Cdc42 or Rac along with Myc-tagged wild-type cortactin. After iso- or hypertonic treatment, the cells were lysed, and the Myc-tagged proteins were immunoprecipitated. Western blot analysis of these precipitates revealed that DN Cdc42 or Rac caused only a modest reduction in the hypertonicity-induced cortactin phosphorylation (Fig. 10B). We therefore conclude that osmotic tyrosine phosphorylation of cortactin does not require the activation of these small GTPases but may be enhanced by it.


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Fig. 10.   Involvement of Rac and Cdc42 in the shrinkage-induced tyrosine phosphorylation of cortactin. A: CHO cells were treated with or without 100 pg/ml of Clostridium difficile toxin B (ToxB) added directly to the growth medium. When complete cell rounding was achieved in the ToxB-treated cells (usually 4 h), the medium was removed and the cells were washed and incubated with Iso or Hyper solutions that contained the same ToxB concentration for an additional 10 min. After cell lysis, cortactin was immunoprecipitated and the precipitates were subjected to Western analysis using anti-PY before being reprobed with anti-cortactin. B: CHO cells were transfected with 2 µg of Myc-cortactin alone (control) or together with either 4 µg of DN Rac or 4 µg of DN Cdc42 as indicated. Subsequently cells were treated with Iso or Hyper solutions and lysed. Lysates were immunoprecipitated with anti-Myc antibody. Precipitates were subjected to Western blotting and the upper half of the membrane was probed with anti-PY to visualize phosphorylated cortactin, whereas the lower part was probed with anti-Myc (bottom) to detect the expression of the transfected small GTPase. Thereafter the upper halves of the membranes were reprobed for anti-Myc to check for the relative expression of Myc-cortactin.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
NOTE ADDED IN PROOF
REFERENCES

Although osmotic stress and other mechanical challenges are known to affect the cytoskeleton, the molecular details of this reorganization are largely unknown. The present study provides novel structural and functional characterization of the hyperosmotic stress-induced cortical cytoskeleton remodeling. Specifically, we show the involvement of newly recognized key regulators of actin dynamics in this process and identify some of the underlying signaling mechanisms. Our results indicate that hyperosmotic shock facilitates incorporation of G-actin into F-actin at the cell boundary concomitant with the peripheral translocation of cortactin and the Arp2/3 complex. Because these proteins have been shown to play a central role in actin nucleation (17, 48, 51), our combined findings strongly support the notion that at least a part of the new F-actin assembly is due to topically upregulated de novo actin nucleation. Consistent with this view, our immunofluorescence and coimmunoprecipitation experiments suggest that hyperosmotic exposure leads to the formation of new nucleation-competent macromolecular complexes that contain Arp2/3, cortactin, and F-actin. This process is independent of the shrinkage-induced tyrosine phosphorylation but appears to be linked to the osmotic stress-evoked activation of Rac and Cdc42. The most plausible sequence of events is the following: osmotic stress stimulates Rac and Cdc42, which in turn activate the Arp2/3 complex and initiate actin nucleation; cortactin is then rapidly recruited to the new nucleation sites where it can further increase filament assembly and stabilize the freshly generated cortical actin fibers. Many of our findings are consistent with this interpretation, but additional mechanisms are also likely to be involved. Here we will discuss the elements of this model and their potential importance in the volume-dependent reorganization of the cytoskeleton.

Increasing evidence supports the notion that various Rho family G proteins may be important upstream mediators in osmotic signaling. Specifically, hypotonicity was suggested to affect ion transport through Rho-dependent pathways (11, 47), whereas our studies provide direct evidence that hyperosmotic stress stimulates Rac and Cdc42 and that this process contributes to volume-dependent cytoskeleton remodeling. Indeed, hypertonic activation of Rac and Cdc42 is consistent with and probably the basis of major signaling events known to be induced by hypertonic stimulation. For instance, hyperosmotic shock has been shown to increase the activity of PAK (8, 42), a common downstream effector of Cdc42 and Rac, as well as Ack, a Cdc42-activatable tyrosine kinase (43). Moreover, osmotic stress caused the translocation of both gamma -PAK and Cdc42 from a soluble to a particulate fraction that included the plasma membrane (42). Importantly, the activation of Rac/Cdc42 and the subsequent PAK stimulation may be at least partially responsible for the observed remodeling events, because the active forms of these proteins have been documented to cause 1) disassembly of stress fibers (2, 31), 2) deposition of peripheral F-actin (for a review see Ref. 4), and 3) translocation of cortactin to the cell border (52). Participation of this mechanism is further supported by our findings that DN Rac and Cdc42 mitigated the osmotic reorganization of F-actin and strongly reduced the translocation of cortactin and Arp3. This result is consistent with the recent finding that DN Rac inhibited the platelet-derived growth factor-induced cortactin accumulation at the periphery (52).

The critical parameters (i.e., cell volume, tonicity, intracellular ion concentrations) and the upstream signaling mechanisms whereby hyperosmotic stress activates Rac and Cdc42 remain to be elucidated. Similarly, future studies should clarify whether changes in the cytoskeletal structure can alter small G protein activity, thereby constituting a feedback loop in mechanochemical signaling.

Active Rac and Cdc42 are known to stimulate the Arp2/3 complex via Wiskott-Aldrich syndrome protein (WASP) and WASP-family verprolin protein (WAVE), respectively (reviewed in Refs. 17, 46). The activated Arp2/3 complex serves as a strong actin-nucleating factor. An initial deposition of F-actin and/or Arp2/3 may be a key event for cortactin recruitment. This view is supported by our findings that the NH2-terminal (Arp2/3- and F-actin-binding) half of cortactin is necessary and sufficient for its osmotic translocation. Recently, Weed et al. (53) reported that both the Arp2/3-binding NTA domain and the F-actin-binding R region are required for the active Rac-induced translocation of cortactin into the ruffles. Although the osmotic redistribution was also more efficient when both domains were present, we observed that the individual domains still showed some peripheral accumulation. This is in agreement with the results of Uruno et al. (48), who reported that the Delta 1-68 mutant lacking the Arp2/3 binding site can still localize to the periphery, and the isolated NTA alone (amino acids 1-80) can colocalize with Arp2/3.

Cortactin is a sensitive and selective marker for the dynamic actin skeleton (54), and its accumulation is a strong indicator of localized de novo F-actin assembly upon osmotic stress. Peripheral cortactin deposition is likely to have important functional implications, because cortactin potentiates actin nucleation and branching and stabilizes the newly assembled actin structure (36, 51). We suggest that the peripheral F-actin accumulation may proceed in two phases: first, an initial WASP/WAVE-induced, Arp2/3-mediated nucleation occurs. This then leads to the recruitment of cortactin, which promotes further polymerization and branching. In essence, cortactin translocation may exert a positive feedback on the formation and stabilization of the peripheral actin network. These processes may reinforce the cell cortex and may represent an important osmoprotective response, i.e., the mammalian counterpart of the cortical remodeling described for Dictyostelium (1, 27, 40, 57).

Cortactin may also have a role in recruiting other components to the periphery, including perhaps the small GTPases themselves. Weed et al. (52) showed that short-term expression of active Rac induces cortactin accumulation under the membrane without an obvious accumulation of Rac itself. This indicates that Rac can "send" cortactin to the periphery without itself going there. During longer-term expression, however, we found strong colocalization between active Rac and cortactin in the ruffles. This finding is consistent with earlier reports showing that active Rac tends to concentrate in ruffles and lamellipodia (33). Moreover, endogenous Rac and cortactin colocalize at the periphery after an osmotic challenge, and we found that Rac translocation was even more pronounced in cortactin-overexpressing cells. This raises the possibility that cortactin might facilitate the recruitment of Rac. Such an effect may not be due to direct binding of active Rac to cortactin, because we have not been able to coprecipitate these molecules (not shown). Rather, this could be a manifestation of a cortactin-induced stabilization of peripheral actin complexes. An enhanced recruitment or retention of Rac in the ruffles could be a novel regulatory way to control local actin dynamics. In agreement with Weed et al. (53), we found that the applied cortactin mutants did not interfere with the localization of endogenous cortactin. Thus new tools are necessary to further assess the various potential roles of cortactin in situ.

Although our results suggest that the Rac/Cdc42 pathway plays a major role in osmotic remodeling of the cytoskeleton, additional mechanisms are also likely to be involved. This notion has been raised by the observation that DN Rac and Cdc42 did not completely abolish F-actin reorganization and cortactin translocation. An explanation for this may be insufficient DN action in some cells. However, participation of Rho family-independent mechanisms is suggested by our recent results obtained in neutrophils. In these cells, hyperosmotic stress caused an approximately twofold increase in total F-actin (41), and ToxB reduced this effect by 60% (28). The remaining component may be due to the shrinkage-induced rise in intracellular ionic force and the decreased hydration state of proteins, both of which were shown to directly augment actin polymerization in vitro (13). Additionally, an osmotically induced change in inositol lipids (9, 49) may promote local actin polymerization and may directly stimulate cortactin deposition, because the fourth repeat of cortactin binds phosphatidylinositol 4,5-bisphosphate (15).

Finally, we investigated the relationship between cortactin tyrosine phosphorylation and translocation. Our results show that these phenomena are not strictly coupled, and the translocation does not require tyrosine phosphorylation. Additionally, although the inhibition of Rac and Cdc42 decreases cortactin tyrosine phosphorylation, Rac and Cdc42 activity is not a prerequisite for this process. Tyrosine phosphorylation has been reported to reduce the actin cross-linking activity of cortactin in vitro (19), and our results suggest that it weakens the association between cortactin and Arp3 in vivo. Future studies should define whether phosphorylation affects the direct interaction of these proteins and/or the association of these proteins with common cytoskeletal components. In either case, tyrosine phosphorylation may be a compensatory process that facilitates the disassembly of the Arp2/3-F-actin-cortactin complex. Such disassembly, enhanced by Src kinases, may be important for the dynamic recycling of the molecule during cell movement: as the leading edge is pushed forward, cortactin at the base of the lamellipodium may become phosphorylated and detach from actin. After dephosphorylation, it may be rebuilt into the new front.

In summary, we have shown that a change in cell volume induces characteristic remodeling of the cortical cytoskeleton and have identified one of the important underlying signaling mechanisms. The activation of small GTPases by osmotic stress or other mechanical stimuli may be a general mechanism whereby changes in cell volume or shape can initiate adaptive responses. In addition to the osmoprotective action, this signaling pathway may play a central role in a variety of complex mechanical phenomena such as the shear stress-dependent remodeling of cell-cell contacts in epithelia.


    NOTE ADDED IN PROOF
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
NOTE ADDED IN PROOF
REFERENCES

While this manuscript was in preparation, it was reported that pharmacological inhibition of tyrosine phosphorylation in platelets did not inhibit the translocation of cortactin from a soluble to a Triton X-insoluble fraction (Lopez I, Duprez V, Melle J, Dreyfus F, Levy-Toledano S, and Fontenay-Roupie M. Thrombopoietin stimulates cortactin translocation to the cytoskeleton independently of tyrosine phosphorylation. Biochem J 356: 875-881, 2001).


    ACKNOWLEDGEMENTS

We are indebted to Dr. S. Grinstein for providing access to the confocal microscope and for valuable discussions. We thank Dr. R. A. Cerione for the HA-Cdc42-expressing cell line, Dr. G. Downey for the Myc-tagged Rac and Cdc42 constructs, Dr. G. L. Lukacs for the 5GFP construct, and Dr. C. A. G. McCulloch for critical reading of the manuscript.


    FOOTNOTES

This work was supported by grants from the Canadian Institutes of Health Research (CIHR) and the Natural Sciences and Engineering Research Council of Canada (NSERC) to A. Kapus. A. Kapus is a CIHR scholar. A. Mak is supported by grants from the Ontario Heart and Stroke Foundation and CIHR. K. Szászi is sponsored by a CIHR postdoctoral fellowship.

Address for reprint requests and other correspondence: A. Kapus, Toronto Hospital, Dept. of Surgery, Transplantation Research, Rm. CCRW 2-850, 101 College St., Toronto, Ontario, Canada M5G 1L7 (E-mail: akapus{at}transplantunit.org).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

May 8, 2002;10.1152/ajpcell.00018.2002

Received 12 January 2002; accepted in final form 6 May 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
NOTE ADDED IN PROOF
REFERENCES

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