Department of Surgery, The Toronto General Hospital University Health Network, and University of Toronto, Toronto, Ontario, Canada M5G 1L7
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ABSTRACT |
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Hypertonicity inhibits a variety of neutrophil functions through poorly defined mechanisms. Our earlier studies suggest that osmotically induced actin polymerization and cytoskeleton remodeling is a key component in the hypertonic block of exocytosis and cell movement. To gain insight into the signaling mechanisms underlying the hyperosmotic F-actin response, we investigated whether hypertonicity stimulates Rac and Cdc42 and, if so, whether their activation contributes to the hypertonic rise in F-actin. Using a recently developed pull-down assay that specifically captures the active forms of these small GTPases, we found that hypertonicity caused an ~2.5- and ~7.2-fold activation of Rac and Cdc42, respectively. This response was rapid and sustained. Small GTPase activation was not mediated by the osmotic stimulation of Src kinases, heterotrimeric G proteins, or phosphatidylinositol 3-kinase. Interestingly, an increase in intracellular ionic strength was sufficient to activate Rac even in the absence of cell shrinkage. Inhibition of Rac and Cdc42 by Clostridium difficile toxin B substantially reduced but did not abolish the hypertonicity-induced F-actin response. Thus hypertonicity is a potent activator of Rac and Cdc42, and this effect seems to play an important but not exclusive role in the hyperosmolarity-triggered cytoskeleton remodeling.
actin cytoskeleton; Rho family GTPases; shrinkage; cell volume
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INTRODUCTION |
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NEUTROPHILS play an indispensable role in host defense against invading microorganisms, but their exuberant activation is a significant contributing factor to local tissue injury (11). Because neutrophil-mediated organ damage represents an important clinical problem, much effort has been focused to find strategies whereby neutrophil functions can be safely and reversibly inhibited. Recent studies from this and other laboratories have shown that hyperosmotic salt solutions exert strong anti-inflammatory effects in various animal models of neutrophil-mediated disease states such as acute respiratory distress syndrome and ischemia-reperfusion injury (2, 3, 33, 34). Resuscitation with hyperosmotic fluids has been shown to reduce neutrophil sequestration in the lung after hemorrhagic shock and lipopolysaccharide-treatment, suggesting that the beneficial immunomodulatory effects may be primarily due to altered neutrophil functions (3, 34). Consistent with this notion, hypertonicity has been reported to abrogate a variety of neutrophil functions including agonist-induced superoxide generation (31), transmigration (38), phagocytosis (21), and degranulation (29, 36).
Although the mechanisms underlying these diverse effects have not been elucidated, our recent studies suggest that an osmotically triggered remodeling of the actin skeleton may play a central role in the observed functional alterations (36). In promyelocytic leukemia (HL-60) cells and normal neutrophils, hyperosmotic stress has been shown to provoke a rapid, sustained, and sizable (~2-fold) increase in F-actin content (19, 20, 36), accompanied by a dramatic spatial reorganization of the cytoskeleton. The newly polymerized F-actin localizes predominantly under the plasma membrane in the form of a thick ring (19, 36) that persists as long as hypertonicity is maintained. This rigid cytoskeletal structure is expected to interfere with functions that require a highly dynamic actin network (e.g., migration and phagocytosis). Moreover, the submembranous actin ring may represent a physical barrier that hinders membrane traffic. Indeed, we found that osmotic shock abolished the exocytosis of all four types of neutrophil granules, and this effect was almost entirely prevented by the pharmacological inhibition of the hypertonically induced actin polymerization (36).
The cellular mechanisms underlying the osmotically triggered cytoskeletal restructuring have remained largely unresolved. In neutrophils, as in other cells, shrinkage stimulates various Src family kinases (27, 28, 30, 35) and leads to the Src family-independent activation of certain mitogen-activated protein kinases, predominantly p38 (25, 35). However, neither of these pathways appeared to play a significant role in the osmotic F-actin response (36), pointing to the participation of alternative signaling cascades. One candidate mechanism might be the osmotic activation of Rho family small GTPases, Rac and/or Cdc42. Several findings give credence to this hypothesis: 1) these proteins are key organizers of the actin skeleton, and as such, they have been implicated in the control of neutrophil locomotion and shape changes (14, 40); 2) addition of activated Cdc42 or Rac has been shown to induce actin polymerization in neutrophil lysates (45) and permeabilized neutrophils (18); and 3) osmotic stress has been found to activate certain downstream effectors of Rac and/or Cdc42, including p21-activated kinase (PAK) and the tyrosine kinase Ack (8, 39).
The aim of the present work was to establish whether hypertonicity activates Rac and Cdc42 and, if so, whether this process participates in the osmotically induced actin polymerization. To address these questions, we used a recently developed pull-down assay (1, 5, 17) that specifically captures the active (GTP bound) forms of these small G proteins, allowing direct determination of their activated states, and Clostridium difficile toxin B, a specific inhibitor of the Rho family. Our results show that hypertonicity causes a strong and sustained stimulation of both small GTPases, and this effect seems to be an important contributor to the osmotic F-actin response. To our knowledge, our studies are the first that directly demonstrate the activation of Rho family GTPases by osmotic stress and that couple this phenomenon to the hypertonicity-induced remodeling of the cytoskeleton.
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MATERIALS AND METHODS |
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Materials.
Dextran, nystatin, pertussis toxin (PTX), wortmannin,
N-formyl-methionyl-leucyl-phenylalanine (fMLP), phorbol
12-myristate 13-acetate (PMA), guanosine
5'-O-(3-thiotriphosphate) (GTPS), GDP, aprotinin,
leupeptin, pepstatin, phenylmethylsulfonyl fluoride (PMSF), and bovine
serum albumin were purchased from Sigma.
4-Amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2) was from Calbiochem. Ficoll, glutathione-Sepharose beads, and
horseradish peroxidase-coupled anti-mouse and anti-rabbit IgG
antibodies were from Amersham. E-lyse was obtained from Cardinal, and
paraformaldehyde was from Canemco. Clostridium difficile
toxin B was purchased from TechLab, and 50X protease inhibitor
cocktail was from PharMingen. Acrylamide was from Bio-Rad, and
nitrocellulose membranes were from Schleicher & Schuell.
Rhodamine-phalloidin was purchased from Molecular Probes. The following
primary antibodies were used: monoclonal anti-phosphotyrosine and
anti-Rac from Upstate Biotechnology; polyclonal anti-Cdc42 from Santa
Cruz Biotechnology; and polyclonal anti-p38 and anti-phospho-p38 from
New England Biolabs.
Neutrophil isolation, media, and cell treatment. Neutrophils were isolated from healthy human volunteers by sedimentation over 3% dextran followed by Ficoll gradient centrifugation (35). Red cells were removed by NH4Cl lysis. Neutrophils were resuspended at the desired concentration in isotonic medium (Iso-Na; osmolarity 295 ± 5 mosmol/l) containing (in mM) 140 NaCl, 3 KCl, 5 glucose, 1 MgCl2, 1 CaCl2, and 10 HEPES (pH 7.4). For permeabilization of the cell membrane of the neutrophil with respect to monovalent ions, the cells were suspended in Iso-K medium containing (in mM) 140 KCl, 10 NaCl, 1 MgCl2, 1 EGTA, 0.194 CaCl2, 5 glucose, and 10 HEPES (pH 7.2), supplemented with 400 U/ml of nystatin and 60 mM sucrose for 5 min. This medium has been shown to maintain the normal isotonic cell volume (30, 36, 41) because the presence of 60 mM sucrose counterbalances the colloid osmotic pressure of intracellular proteins, the predominant driving force that dictates fluid distribution in cells permeable to monovalent ions. In certain experiments, intact neutrophils were pretreated with various inhibitors. The following agents and preincubation times were used: PTX: 570 ng/ml, 1 h; C. difficile toxin B: 90 µg/ml, 1.5 h; wortmannin: 100 nM, 30 min; and PP2: 10 µM, 10 min. Hypertonicity was induced by increasing the osmotic concentration of the media by 200 mosmol/l added in the form of 100 mM NaCl, 200 mM sucrose, or 100 mM KCl, as specified for the corresponding experiments. For chemical stimulation of neutrophils, either fMLP (100 nM or 1 µM, 30 s to 1 min) or PMA (100 nM for 10 min) was used.
Glutathione-S-transferase-p21-binding domain bead preparation. A pGEX-2T vector with an insert consisting of amino acids 67-150 of PAK [the p21-binding domain (PBD) of PAK] fused to glutathione-S-transferase (GST) was kindly provided by Dr. G. Bokoch (Scripps Institute, La Jolla CA). This vector was used to transform Escherichia coli that were induced to produce the protein product using 1 mM isopropylthioglucose. After disruption of the bacterial cells by sonication in the presence of 10 µg/ml aprotinin, 10 µg/ml leupeptin, 10 µg/ml pepstatin, and 1 mM PMSF, the bacterial lysate was incubated for 1 h with glutathione-Sepharose beads. The beads were then washed four times with phosphate-buffered saline supplemented with 5 mM EDTA.
Rac and Cdc42 activation assays.
The assay was performed as reported earlier (1, 5) with
small modifications. Briefly, 107 neutrophils in a volume
of 700 µl per experimental condition were prepared and treated with
various inhibitors as described above. Cells were exposed to the
desired stimulus and then lysed in an equal volume of 2× lysis buffer
containing 20 mM HEPES (pH 7.4), 150 mM NaCl, 2% Nonidet P-40 (NP-40),
20% glycerol, 8 mM EGTA, 8 mM EDTA, and 2× protease inhibitor
cocktail. In agreement with previous reports (1, 5), this
direct cell lysis method provided the most consistent and reproducible
results. The lysates were cleared with a brief centrifugation at 12,000 rpm and then incubated in the presence of GST-PBD beads for 30 min with
constant rotation at 4°C. Subsequently, the beads were washed twice
in a buffer containing 20 mM HEPES (pH 7.4), 142.5 mM NaCl, 1% NP-40, 10% glycerol, 4 mM EGTA, and 4 mM EDTA. To obtain positive and negative controls for the active and inactive small GTPases, the nonhydrolyzable GTP analog GTPS and GDP were used, respectively. Cell lysates obtained from untreated cells were incubated with 100 µM
GTP
S or 1 mM GDP in the presence of 10 mM EDTA for 15 min at 30°C
to ensure efficient loading with the added nucleotide. To terminate the
reaction and to "lock in" the nucleotides, the lysates were placed
on ice and supplemented with 60 mM MgCl2. These control
samples were then incubated with the GST-PBD beads and washed in the
same manner as the other samples. Captured proteins were removed from
the beads by boiling the samples in Laemmli buffer, and samples were
subjected to SDS-PAGE and Western blotting.
SDS-PAGE and Western blotting. Proteins captured by the beads or aliquots from whole cell lysates were separated on 10% or 15% polyacrylamide gels, as specified, and then transferred to nitrocellulose membranes with a Bio-Rad Mini Protean II apparatus. Membranes were blocked in Tris-buffered saline containing 5% bovine serum albumin and then incubated with the corresponding primary antibodies for 1 h. After thorough washing, the membranes were incubated with horseradish peroxidase-conjugated anti-mouse or anti-rabbit secondary antibodies for 1 h. The blots were repeatedly washed, and the immunoreactive bands were visualized using the enhanced chemiluminescence system. Densitometric analysis on blots was performed using a Bio-Rad model GS-690 imaging densitometer and Molecular Analyst software (version 1.5) as described (28).
Superoxide production. Cells pretreated with or without various inhibitors were suspended in Iso-Na medium containing 100 µM ferricytochrome c and either left untreated or challenged with 1 µM fMLP for 5 min. Cells were then sedimented with a brief spin, and the superoxide-induced cytochrome reduction was determined by measuring the increase in absorbance at 550 nm.
F-actin quantification and flow cytometry. Neutrophils were suspended in Iso-Na medium at a concentration of 1 × 106 cells/ml and then challenged with either fMLP (100 nM for 1 min) or hypertonicity (100 mM NaCl for 10 min). Cells were then fixed with paraformaldehyde (final concentration 4%) for 15 min, pelleted, and resuspended in 100 µl isotonic buffer containing 0.1% Triton X-100 and 0.33 µM rhodamine-phalloidin for 15 min at 4°C. Stained cells were pelleted and washed twice in isotonic medium before flow cytometric analysis as described previously (36). Typically, 5,000 cells per condition were measured with a Beckman Coulter flow cytometer (model Epics XL/MCL; excitation 488 nm, emission 575 nm) driven by System II software. The results are expressed as percent change compared with the isotonic control.
Data are presented as means ± SE for the number of experiments indicated (n) or as representative immunoblots for at least three similar experiments. Statistically significant (P < 0.05) difference among the treatment groups was calculated using Student's t-test. ![]() |
RESULTS |
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Osmotic stress activates the small G proteins Rac and Cdc42.
To assess whether hypertonicity affects the activity of Rho family
small G proteins, we compared the amount of activated Rac and Cdc42 in
lysates obtained from isotonically or hypertonically treated
neutrophils. The PBD-GST beads precipitated only marginal amounts of
Rac and Cdc42 from isotonically treated cells (Fig. 1A, Iso). In contrast,
hyperosmotic exposure (100 mM NaCl added into the isotonic medium for
10 min) caused a strong increase in the amount of both active Rac and
Cdc42 [Fig. 1A, Hyp 10' (NaCl)]. Similar activation was
observed when osmolarity was raised to the same level using 200 mM
sucrose, a nonionic osmolyte [Fig. 1A, Hyp 10' (Sucrose)].
This latter finding excluded the possibility that the enhanced
association of the small GTPases to the beads was due to the higher
salt concentration present in the lysates of the NaCl-treated cells.
Thus the assay reflected the activation of the small G proteins before
cell lysis, and this effect was independent of the type of the
shrinkage-inducing compound.
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Osmotic activation of Rac and Cdc42 does not require Src kinase-mediated tyrosine phosphorylation and appears to be independent of heterotrimeric G proteins and phosphatidylinositol 3-kinase. The upstream signaling mechanisms whereby chemoattractants stimulate the Rho family small GTPases are incompletely understood. Although the literature contains several controversies, three major pathways have been implicated: protein tyrosine phosphorylation (5, 6, 9), activation of heterotrimeric G proteins (1, 17, 23), and stimulation of phosphatidylinositol (PI) 3-kinase (1, 5, 22). Because hyperosmotic shock has been suggested to target each of these pathways (13, 28, 35, 43), we investigated their potential involvement in the osmotically triggered Rac/Cdc42 response.
Cell shrinkage induces a robust tyrosine phosphorylation in a number of neutrophil proteins (Fig. 3A and Refs. 30 and 35). Pretreatment of the cells with the Src-family inhibitor PP2 completely prevented the osmotically induced phosphotyrosine accumulation (Fig. 3A). In contrast, PP2 failed to significantly affect the osmotic activation of Rac (Fig. 3B): densitometric analyses of five blots resulting from such experiments showed that hyperosmolarity caused a 1.65 ± 0.3- and 1.7 ± 0.3-fold (n = 5) stimulation in the absence and presence of PP2, respectively.
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An increase in intracellular ionic strength is sufficient to
activate Rac.
An increase in the extracellular concentration of impermeable osmolytes
results in an instantaneous decrease in cell volume and a parallel
increase in the intracellular solute concentrations. Intriguingly, our
previous studies have suggested that the critical parameter in the
hypertonicity-induced actin response may be the rise in the salt
concentration rather than the reduction in the cell volume per se
(36). To address whether the hypertonic activation of
small GTPases might also be related to elevation of intracellular ion
concentrations, we used nystatin-permeabilized cells in which the
intracellular ion concentrations can be manipulated independently of
the cell volume (30, 36, 41). The colloid osmotic pressure was balanced by 60 mM sucrose. In this experimental system the cells
keep near-normal volume that does not change upon addition of
monovalent ions into the medium (30, 36). Because the
membrane of nystatin-treated cells is freely permeable to monovalent
ions, we used a KCl-based medium that is similar to the normal
intracellular milieu. Replacing the isotonic NaCl medium with an
isotonic KCl medium had no effect on Rac (Fig.
4, lanes 1 and 3).
Moreover, addition of an extra 100 mM NaCl to nonpermeabilized cells
resulted in an equal Rac activation regardless of whether the cells
were suspended in isotonic NaCl or KCl medium (Fig. 4, lanes
2 and 4), indicating that extracellular KCl per se does
not influence the response. Addition of nystatin alone had no
significant effect on Rac (Fig. 4, lane 5). Importantly,
addition of extra NaCl to nystatin-permeabilized cells resulted in a
strong Rac activation (Fig. 4, lane 6). Addition of KCl also
provoked a significant (albeit somewhat smaller) response, implying
that the activation is not Na specific (Fig. 4, lane 7).
These observations suggest that an increase in the concentration of
intracellular monovalent ions (or in the overall ionic strength) is
sufficient to activate Rac without a concomitant reduction in the cell
volume. Furthermore, both the small GTPase activation and the F-actin
response seem to be sensitive to the same parameter.
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Small G protein activation contributes to the hypertonically
induced actin polymerization.
To test whether the hypertonic activation of small GTPases could be
responsible for the osmotically induced increase in F-actin, we used
C. difficile toxin B, an enzyme that inactivates the Rho family small G proteins by glycosylating them at a critical threonine residue (26). Because neutrophils are much less sensitive
to this toxin than fibroblasts (40), we first performed
controls to assess whether toxin B effectively inhibited the small
GTPases under our conditions. Figure
5A shows that preincubation of
neutrophil for 2 h with 90 µg/ml toxin B almost completely
abolished the hypertonic activation of Rac. To verify efficiency with
an independent method, we followed the effect of toxin B on p38
activation, a process thought to be downstream of small GTPases in
various cell types. As shown in Fig. 5B, toxin B prevented
the hyperosmolarity-induced phosphorylation of p38. Additionally, toxin
B almost completely abolished the fMLP-triggered activation of p38 as
well. The total p38-labeling was the same in the absence and presence
of toxin B, indicating that no loss of kinase occurred from the toxin
B-treated neutrophils (Fig. 5B, bottom). Thus we
can conclude from these experiments that toxin B efficiently permeated
the neutrophils and inhibited the small GTPases. Moreover, the Rho
family small GTPases appear to be involved in the activation of p38 by
hypertonicity and fMLP.
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DISCUSSION |
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The two major findings of the present study are that hypertonicity activates Rac and Cdc42 in neutrophils and that the activation of these small GTPases contributes to the hyperosmolarity-induced actin polymerization. To our knowledge, our studies are the first to provide direct evidence for the osmotic stimulation of these small GTPases. This phenomenon may be a central upstream event responsible for a variety of known biochemical effects of hyperosmolarity. For example, it may underlie the osmotic stimulation of the serine/threonine kinase PAK (8) or the tyrosine kinase Ack (39), as these enzymes are direct downstream effectors of Rac or Cdc42. Indeed, our inhibitor studies suggest that small GTPases act as upstream mediators in the hyperosmotic activation of p38 in neutrophils, a finding consistent with the well-known role of the Rac/Cdc42-PAK pathway in p38 regulation (32, 44). Nevertheless, it is worth noting that the involvement of small GTPases in the hypertonic activation of p38 appears to differ in various cell types. In a recent paper, Clerk et al. (7) reported that toxin B failed to prevent the sorbitol-induced p38 phosphorylation in myocytes, although it effectively inhibited c-Jun kinase phosphorylation. In agreement with this, we found that the osmotically induced p38 activation was not sensitive to toxin B in Chinese hamster ovary cells (unpublished observation).
Our ongoing studies imply that activation of the Rho family GTPases upon osmotic stress is not restricted to neutrophils but also occurs in attached cells such as fibroblasts, which contain a more complex cytoskeleton. Consistent with this notion, Roig et al. (37) recently reported that in NIH/3T3 cells, hyperosmolarity induced the translocation of Cdc42 from a soluble to a particulate fraction, a phenomenon suggestive of Cdc42 activation. These data raise the possibility that the Rho family GTPases may play a general role in mechanochemical signal transduction as upstream mediators that relay the effects of osmotic stress and other mechanical stimuli to various stress kinases and the cytoskeleton.
The mechanisms whereby hyperosmolarity and other stimuli regulate Rac and Cdc42 are incompletely understood. The control of Rho GTPases is an extremely complex process involving various guanine nucleotide exchange factors (GEFs), GDP dissociation inhibitors (GDIs), and GTPase-activating proteins (GAPs) (for review, see Ref. 42). Increased GTP-binding, and thereby enhanced small G protein activity, may be brought about by activation of GEFs and/or the inhibition of GDIs and GAPs. Vav is an important GEF highly expressed in hematopoietic cells that has been shown to be activated by Src family-dependent tyrosine phosphorylation (6, 9) and lipid products of PI 3-kinase (22, 42). Because hyperosmolarity had been shown to affect these pathways, their involvement in the mediation of the osmotic effects was an attractive hypothesis. However, our pharmacological data do not support their role in the osmotic Rac activation. Consistent with this finding, Src family and PI 3-kinase-independent Rac activation has been observed in fMLP and PMA-stimulated neutrophils as well (1, 17). The role of PI 3-kinase in the fMLP-triggered response remains controversial because two groups found that wortmannin reduced or prevented small GTPase activation (1, 5), whereas another team observed no effect (17). These conflicting results may partly originate from the existence of different Rac isoforms (Rac1 and 2), the regulation of which may be distinct. Future studies should define whether the osmotic sensitivity of Rac is isoform specific. Another candidate mechanism was the activation of heterotrimeric G proteins, which have been implicated in the hyperosmotic stimulation of Na+/H+ exchange (12) and stress kinases (13) and are thought to be direct activators of certain GEFs (17, 23). However, we found no evidence for the requirement of PTX-sensitive G proteins in the osmotic effect. While the exact mechanism of small GTPase activation remains to be clarified, we have made the intriguing observation that a rise in intracellular salt concentration was sufficient to activate Rac. This raises the possibility that increased intracellular salinity may directly or indirectly promote GEF or reduce GAP activity, or help dissociate GDIs from Rac.1 The overall increase in cytosolic salt concentration achieved after shrinking intact cells in a 500 mosmol/l medium is comparable to that achieved by adding an extra 100 mM salt to permeabilized neutrophils. Nevertheless, the contribution of increased intracellular ionic strength to the shrinkage-induced Rac activation in intact neutrophils requires further studies, and our results by no means exclude an additional role for volume reduction per se.
Irrespective of the underlying mechanisms, the question arises whether the activation of small GTPases contributes to the actin response. Several observations are consistent with or directly support this possibility. First, the onset and the time course of the small GTPase response are compatible with such an effect. Interestingly, hyperosmolarity induces a sustained elevation both in small GTPase activity and the F-actin level, whereas fMLP causes only a transient small GTPase activation accompanied by a transient rise in F-actin. Second, both Rac activation and the rise in F-actin can be elicited by elevated salt concentration. Finally, and most importantly, prevention of small GTPase activation by C. difficile toxin B significantly reduces the osmotically induced actin response.
In vivo, actin polymerization may occur through three mechanisms: de novo nucleation, severing of actin filaments, and dissociation of capping proteins from filament ends (10). The latter two mechanisms generate new free ends for polymerization. Recent breakthrough discoveries have revealed the link between small GTPases and de novo actin nucleation (reviewed in Ref. 24). In a variety of cell types, active Cdc42 and Rac stimulate WASP and Wave proteins, respectively, which in turn activate the actin-related protein (Arp2/3) complex, a central nucleation-inducing factor. In neutrophils, Rac appears to promote F-actin formation through an Arp2/3-independent pathway (18), presumably by inducing dissociation of the capping protein gelsolin (4). These mechanisms may all participate in the hyperosmotic, small GTPase-mediated F-actin formation.
Our results show that a sizable part (40-50%) of the osmotic actin response is preserved in the presence of toxin B. One explanation could be that the inhibition of small GTPases by toxin B was incomplete. Although this possibility cannot be excluded at present, it seems less likely given the facts that toxin B abolished the osmotic p38 activation and reduced the fMLP-triggered actin rise by >80% (not shown). Moreover, we found that expression of dominant negative Rac and Cdc42 in fibroblasts strongly reduced but did not abolish the shrinkage-induced reorganization of F-actin and actin-binding proteins (unpublished observations). We therefore favor the interpretation that the actin response contains a small GTPase-independent component(s). Such factors may include shrinkage-related physicochemical changes, e.g., osmotic dehydration of actin filaments or increased ionic strength, both of which have been reported to promote F-actin assembly in vitro (16). These factors also may alter the activity of capping and severing proteins. Additionally, other volume-dependent signals, such as changes in membrane phosphoinositides (43), also may influence actin polymerization in a small GTPase-independent manner.
In summary, our studies provide evidence that hypertonicity activates two Rho family small GTPases and suggest that this event is one of the mechanisms whereby osmotic stress induces actin polymerization. The ensuing cytoskeleton remodeling may represent an important osmoprotective response that reinforces the cell cortex and, at the same time, may be one of the major mechanisms underlying the neutrophil-inhibitory effects of hyperosmotic stress.
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ACKNOWLEDGEMENTS |
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We are indebted to Drs. Katalin Szászi, Sandro Rizoli, and Sergio Grinstein for valuable discussions.
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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 (to A. Kapus). A. Kapus is a CIHR scholar. A. Lewis is a recipient of a research training fellowship from Dalhousie University.
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).
1
It is worth noting here that high salinity and
hypertonicity were found to increase the transcription of
1-chimerin, a neuronal specific Rac-GAP
(15). This phenomenon may represent a long-term negative
feedback mechanism that downregulates the osmotically stimulated small GTPases.
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.
First published October 10, 2001; 10.1152/ajpcell.00427.2001
Received 6 September 2001; accepted in final form 11 October 2001.
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