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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ABSTRACT |
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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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INTRODUCTION |
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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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MATERIALS AND METHODS |
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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 -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
DH5, 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
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 (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 GTP
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). ![]() |
RESULTS |
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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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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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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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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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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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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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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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DISCUSSION |
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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 -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 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.
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NOTE ADDED IN PROOF |
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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).
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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