Department of Surgery, Toronto General Hospital, and the University of Toronto, Toronto, Ontario, Canada M5G 1L7
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ABSTRACT |
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Hypertonicity suppresses neutrophil functions by unknown mechanisms. We investigated whether osmotically induced cytoskeletal changes might be related to the hypertonic inhibition of exocytosis. Hyperosmolarity abrogated the mobilization of all four granule types induced by diverse stimuli, suggesting that it blocks the process of exocytosis itself rather than individual signaling pathways. Concomitantly, osmotic stress provoked a twofold increase in F-actin, induced the formation of a submembranous F-actin ring, and abolished depolymerization that normally follows agonist-induced actin assembly. Several observations suggest a causal relationship between actin polymerization and inhibition of exocytosis: 1) prestimulus actin levels were inversely proportional to the stimulus-induced degranulation, 2) latrunculin B (LB) prevented the osmotic actin response and restored exocytosis, and 3) actin polymerization induced by jasplakinolide inhibited exocytosis under isotonic conditions. The shrinkage-induced tyrosine phosphorylation and the activation of the Na+/H+ exchanger were not affected by LB. Inhibition of osmosensitive kinases failed to prevent the F-actin change, suggesting that the osmotic tyrosine phosphorylation and actin polymerization are independent phenomena. Thus cytoskeletal remodeling appears to be a key component in the neutrophil-suppressive, anti-inflammatory effects of hypertonicity.
cytoskeleton; shrinkage; osmotic shock; tyrosine kinases; latrunculin B
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INTRODUCTION |
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NEUTROPHILS PLAY A PIVOTAL ROLE in antimicrobial host defense, however, their uncontrolled activation is a key pathological factor in a variety of inflammatory conditions, such as ischemia-reperfusion injury and the adult respiratory distress syndrome. Recent studies from this and other laboratories have shown that hyperosmotic solutions can exert strong anti-inflammatory effects in vivo, primarily by interfering with neutrophil transmigration into the tissues (22, 37). Hypertonicity has been shown to inhibit a plethora of neutrophil functions, including superoxide production (26), release of certain granular contents (24), expression of surface attachment molecules (37, 38), phagocytosis, and migration (39, 46). Although these effects are well documented, the underlying cellular mechanisms have remained largely undefined.
To find a link between volume change and the ensuing functional consequences, increasing attention has been focused to identify signaling pathways activated by osmotic shock. One of the most prominent responses to cell shrinkage is a dramatic increase in protein tyrosine phosphorylation (23, 25, 38, 44). We and others have shown that shrinkage results in the activation of at least two kinase pathways in neutrophils (22, 25, 38). It leads to the stimulation of members of the Src family (e.g., Hck), which in turn propagate the signal to other nonreceptor tyrosine kinases (e.g., Syk, Pyk2), and it also initiates the Src family-independent activation of certain mitogen-activated protein kinases, especially p38 (23, 38). The relationship between these kinase cascades and the alteration in the various functions has not been elucidated. So far, the only functional consequence that has been conclusively associated with hypertonicity-induced kinase activation is the p38-dependent cleavage of L-selectin from the neutrophil surface (38). The attempt to correlate the activation of the different kinase pathways with the inhibition of the above-mentioned neutrophil functions is further complicated by the fact that these cascades are generally activated by most of the stimuli that are strong enhancers of these same neutrophil functions. Thus it is not clear whether the hypertonically treated neutrophils remain suppressed because of, or rather in spite of, the activation of the various kinase cascades.
Another key target of the hypertonic effect might be the cytoskeleton, the integrity of which is a prerequisite for several neutrophil functions. Surprisingly, little is known about the effect of cell shrinkage on the actin skeleton, and the potential correlation between such a change and the osmotic inhibition of neutrophils has not been addressed. Earlier studies dealing with this issue have concentrated on the potential role of actin in regulatory volume changes. Neutrophil swelling was associated with a decrease in filamentous actin (F-actin) content (12), and an inverse relationship between volume and polymerized actin was observed in HL-60 cells (17, 18). On the basis of these facts, we hypothesized that hypertonicity interferes with the normal polymerization/depolymerization cycle and/or distribution of actin in neutrophils, and this effect may be an important common mechanism whereby many neutrophil functions can be altered.
The aim of the present study was to explore the potential role of the hypertonicity-induced actin remodeling in the osmotic alteration of cellular function. We have selected exocytosis as an important functional representative because the actin network is known to affect degranulation (1, 2, 5, 14, 34). In addition, the presence of four different granule types, the exocytosis of which is regulated by a multitude of signaling pathways, offers an excellent opportunity to establish whether hyperosmolarity causes a general block in membrane traffic or whether it selectively interferes with specific signaling routes. We also intended to discern whether the activation of osmosensitive kinases may lay upstream or downstream of the observed cytoskeletal effects.
These studies provide evidence that hypertonicity, independent of tyrosine phosphorylation, markedly alters F-actin content, distribution, and dynamics, and these changes play a crucial role in the osmotic inhibition of exocytosis and other neutrophil functions.
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MATERIALS AND METHODS |
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Materials. Ficoll and dextran T-500 were purchased from Pharmacia Biotech, N-formyl-methionyl-leucyl-phenylalanine (FMLP), phorbol 12-myristate 13-acetate (PMA), Escherichia coli O111: B4 lipopolysaccharide (LPS), cytochalasin B (CB), sucrose, nystatin, bovine serum albumin (BSA), and diisopropyl fluorophosphate (DFP) were from Sigma, Triton X-100 was from Caledon Laboratories, and the enhanced chemiluminescence detection system (ECL), protein G-agarose beads, and horseradish peroxidase-coupled anti-rabbit and anti-mouse antibodies were from Amersham. Formaldehyde was obtained from Canenco. 2',7'-Bis(2-carboxyethyl)-5(6)-carboxyfluorescein-acetoxymethyl ester (BCECF-AM), fura 2-AM, SlowFade antifade kit, rhodamine phalloidin, and jasplakinolide (JK) were from Molecular Probes. Genistein, SB-203580, 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2), A-23187, ionomycin, and latrunculin B (LB) were purchased from Calbiochem. Proteinase inhibitor cocktail 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 in pure ethanol was from PharMingen. All chemicals used were of the highest purity available.
Antibodies. Monoclonal anti-phosphotyrosine (4G10) was purchased from Upstate Biotechnology, and the polyclonal phospho- and nonphospho-anti-p38 were from New England Biolab. Polyclonal anti-actin was from Sigma, monoclonal anti-Syk and polyclonal anti-Hck were from Santa Cruz Biotechnology, and monoclonal anti-Pyk2 was from Transduction Laboratories. We purchased from Serotec the anti-CD63 antibody, the fluorescein isothiocyanate-conjugated (FITC) rat anti-mouse IgM heavy chain antibody, the monoclonal FITC anti-L-selectin, the FITC anti-CD66b, and the phycoerythrin (PE)-labeled anti-CD35. The PE-labeled anti-CD11b was from Becton Dickinson. Neutrophil gelatinase B release was measured with the use of a sandwich ELISA kit from Amersham Pharmacia Biotech.
Neutrophil isolation and media. Cells were isolated by Ficoll gradient from fresh blood drawn by venipuncture as described earlier (37). Isolated neutrophils were suspended in DMEM (GIBCO BRL) supplemented with 10% fetal calf serum (Hyclone Lab) plus penicillin/streptomycin (GIBCO BRL), in bicarbonate-free RPMI 1640 (GIBCO BRL) buffered to pH 7.4 with 10 mM HEPES, or in isotonic saline buffer containing (in mM) 140 NaCl, 5 KCl, 5 glucose, 1 MgCl2, 1 CaCl2, and 10 HEPES (pH 7.4). Hypertonic medium (500 mosmol/kgH2O) was obtained by the addition of an extra 100 mM NaCl. Iso-K medium contained (in mM) 140 KCl, 10 NaCl, 1 MgCl2, 1 EGTA, 0.194 CaCl2 (100 nM free Ca2+), 5 glucose, and 10 HEPES (pH 7.2). To permeabilize the cells for monovalent cations, the Iso-K medium was supplemented with 400 U/ml nystatin and 60 mM sucrose. Sucrose was included to counterbalance the intracellular colloidosmotic pressure and thereby maintain cell volume by preventing swelling of the permeabilized cells as reported earlier (44). When indicated, the nystatin-containing permeabilization buffer was supplemented with 100 mM KCl, 100 mM NaCl, or 200 mM sucrose.
Granule exocytosis. Exocytic insertion of CD35, CD66b, CD63, and CD11b into the cell membrane was determined by flow cytometry. Neutrophils suspended in DMEM at 1 × 106/ml were incubated with various agents under isotonic or hypertonic conditions as detailed in the figures. Subsequently, 100-µl aliquots of cells were incubated at 1:10 dilution of FITC- or PE-labeled antibodies for 20 min at 4°C and analyzed on a FACScan (Becton Dickinson) with the use of an FL1 detector (488-nm argon laser excitation and 520-nm emission wavelengths) and FL2 detector (570-nm emission wavelength), respectively. In all experiments, 5,000 cells were typically analyzed per condition and the results expressed as mean channel fluorescence. Primary granule exocytosis was studied in the presence of CB (10 µM) because otherwise only a negligible release occurs (40). For CD63 determination, after incubation with anti-CD63 antibody, the cells were pelleted and resuspended in isotonic medium with 1:50 dilution of FITC-anti mouse IgM antibody. Exocytic release of gelatinase into the supernatant was quantified by an ELISA commercially available. After treatment, supernatant was collected by centrifugation, and gelatinase release was measured in accordance with the manufacturer's instructions. Neutrophils (1 × 106/ml) suspended in isotonic or hypertonic saline buffer were treated with either FMLP (100 nM) for 20 min or PMA (10 nM) or A-23187 (1 µM) for 10 min at 37°C.
Quantification and distribution of intracellular F-actin. F-actin was quantified essentially as described (19). A 500-µl aliquot containing 1 × 106 neutrophils in either isotonic saline buffer or Iso-K media was incubated at 37°C with various agents under isotonic or hypertonic conditions as detailed in the figures. Cells were then fixed in the corresponding solution containing 4% paraformaldehyde for 15 min and pelleted and resuspended in isotonic buffer supplemented with 0.1% Triton X-100 and 0.33 µM rhodamine phalloidin for 15 min at 4°C. Subsequently, the cells were washed and pelleted again. F-actin content was quantified by flow cytometry (480-nm excitation and 570-nm emission). Five thousand cells were typically measured per condition and the values expressed as percentages of control samples. Occasionally, batches of cells were stained with either nitrobenzoxadiazole-phallacidin or rhodamine phalloidin. The results obtained with these two dyes were indistinguishable. For the investigation of F-actin distribution, fixed and rhodamine phalloidin-labeled cells were centrifuged onto glass coverslips by Cytospin. The slides were mounted with the use of the SlowFade kit and visualized by fluorescent microscopy with the use of a Leica DRBM inverted fluorescence microscope connected to a cooled charge-coupled device video camera (Princeton Instruments). Images were collected with Winview software.
SDS-PAGE and immunoblotting. After specified treatments, the neutrophils were rapidly sedimented in a microcentrifuge at 4°C, and the pellet was resuspended in hot Laemmli sample buffer (LSB, 10% glycerol, 5% 2-mercaptoethanol, 2% SDS, 0.025% bromphenol blue, and 62.5 mM Tris, pH 6.8) and boiled for 10 min. Samples were subjected to 10% SDS-PAGE, transferred to a nitrocellulose membrane, and blocked for 1 h at room temperature in Tris-buffered saline containing 5% BSA. The membranes were then incubated with the corresponding antibodies at room temperature for 1 h. The dilution was 1:3,000 for anti-phosphotyrosine (4G10), 1:200 for anti-actin, and 1:1,000 for anti-p38, anti-Syk, anti-Pyk2, and anti-Hck. After being washed, the membranes were incubated with either peroxidase-conjugated anti-mouse or anti-rabbit secondary antibody (1:4,000) and visualized with the use of ECL.
Immunoprecipitation. To minimize proteolysis, neutrophils were pretreated with 1 mM DFP for 20 min, washed, and resuspended at a concentration of 5 × 106 cells/ml in HEPES-buffered RPMI. After specific treatments, the same volume of ice-cold buffer of the corresponding osmolarity was added, and the cells were rapidly sedimented in a microcentrifuge. The cells were dissolved in ice-cold lysis buffer containing (in mM) 100 NaCl, 30 HEPES, 20 NaF, 1 EGTA, 1 sodium vanadate, 1% Triton X-100, and 20 µl/ml proteinase inhibitor cocktail. The lysate was kept on ice for 10 min, and the Triton-soluble part was obtained by centrifugation (10 min at 4°C), precleared, and incubated with the relevant antibody for 2 h. Subsequently, the lysate was incubated with protein G-agarose beads at 4°C for 1 h. The immune complexes were then sedimented and washed three times with washing buffer (25 mM Tris, pH 7.4, 1 mM sodium vanadate, and 150 mM NaCl), and the precipitate boiled in LSB. For cytoskeleton-associated actin determination, the Triton-insoluble pellet was dissolved and boiled in LSB and then separated by SDS-PAGE.
Measurement of cell volume. Neutrophil volume was determined by electronic sizing with the use of a Coulter counter (model ZM) equipped with a Channelyzer. Typically, 5,000 cells were analyzed per run. Cell distribution among 75 channels was followed, and the volume corresponding to the median channel (~250-300 cells/channel) was determined. Approximately 65% of the cells had their volumes within ± 10% of the median.
Measurement of intracellular pH and intracellular free Ca2+. These parameters were followed fluorometrically (43). Briefly, for intracellular pH determination, 1 × 106 neutrophils suspended in isotonic saline were incubated with 2 µM 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein-AM for 15 min at 37°C. The cells were then washed and resuspended in isotonic saline buffer, and the fluorescence was continuously monitored with the use of a Perkin-Elmer LS50 fluorimeter. The excitation and emission wavelengths were 490 nm and 520 nm, respectively. For intracellular free Ca2+ determination, cells were loaded with 4 µM fura 2-AM for 30 min. Ratio fluorescence was monitored with the use of dual excitation (340/380 nm) and single emission (510 nm).
Electron microscopy. Cells were fixed for 2 h in 2.5% glutaraldehyde in PBS, pelleted and fixed in 1% osmium tetroxide, dehydrated in graded ethanol, and embedded in Epon-Araldite resin. Thin sections were mounted on copper grids and stained with uranyl acetate and lead citrate for viewing in a Philips 400 electron microscope.
Statistical analysis. Data are presented as means ± SE of the means for n experiments as indicated with duplicate readings within each experiment. Blots are representative of at least three independent studies. Significance was assessed with the use of one-way ANOVA with post hoc testing using the Student-Newman-Keuls multiple-comparison test. A probability of P < 0.05 was considered significant.
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RESULTS |
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The effect of hypertonicity on neutrophil granule exocytosis.
The fusion of the various neutrophil granules with the plasma membrane
is triggered by multiple signaling pathways (3, 40). To
assess whether hypertonicity can exert a universal inhibition on the
process of exocytosis per se, or whether its effect is restricted to
certain granule types and/or to particular signaling routes, we exposed
the cells to a variety of stimuli known to act through distinct
mechanisms, and we measured the exocytosis of each of the four granule
types under isotonic and hypertonic conditions. To ascertain
specificity, we used granule markers that were stored exclusively in
one vesicle type and that were inserted into the membrane solely by
exocytosis. Thus to detect the insertion of primary granules,
secondary granules, and secretory vesicles, the surface expression of
CD63, CD66b, and CD35 was followed, respectively. Surface labeling was
detected on nonpermeabilized cells with the use of flow cytometry. To
follow the fusion of tertiary granules that do not contain one
exclusive marker but are the major source of gelatinase B, we measured
the release of this enzyme into the extracellular space by ELISA. The
agents applied to trigger exocytosis included both receptor-mediated and receptor-independent stimuli. Specifically, we used the bacterial peptide FMLP, which requires both Ca2+ signal and the
activation of tyrosine kinases to elicit exocytosis (30);
LPS, with its effect depending on the activation of p38 (37); the protein kinase C activator phorbol ester that
induces granule release in a Ca2+ signal- and tyrosine
kinase-independent manner (28); and the ionophore A-23187,
which acts via the direct elevation of intracellular Ca2+.
Figure 1 shows that under isotonic
conditions, each of these agents was able to induce significant
exocytosis, although their efficiency in the mobilization of the
different vesicles varied, which is in agreement with earlier findings
(3). Each of the four stimuli caused similar and
substantial exocytosis of secretory vesicles and secondary granules,
whereas tertiary granules were best mobilized with PMA, and the primary
granules were most responsive to A-23187 and resistant to LPS,
confirming earlier findings (40). When these stimuli were
added to hyperosmotically shrunken cells, a dramatic reduction in
exocytosis was observed. This phenomenon was not due to altered epitope
recognition by the corresponding antibodies because flow cytometry was
performed after resuspending the cell in the same isotonic solution.
The osmotic prevention of exocytosis was essentially complete and
affected all four granule types, suppressing their mobilization to near
baseline levels.1
Importantly, hypertonicity abrogated exocytosis regardless of the
applied stimulus. Both Ca2+-dependent and -independent
responses were prevented, and even the direct, A-23187-elicited granule
release was essentially abolished, implying that mechanisms other than
the previously suggested hyperosmotic distortion of the
Ca2+ signal (24) must be contributing to the
observed effects. Taken together, our data show that hyperosmotic
exposure of neutrophils results in a universal block of granule
exocytosis. This inhibition is dependent neither on granule type nor on
stimulus, suggesting that the underlying mechanisms may affect a common
step in the exocytic process.
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The effect of hypertonicity on the actin cytoskeleton. The actin skeleton is known to play a crucial role in most neutrophil functions, including exocytosis (20). It is well established that agents that inhibit actin polymerization (e.g., CB) potentiate stimulus-induced degranulation, and polymerized actin has been suggested to act as a barrier against exocytosis in other cell types (1, 2, 5, 14, 34). It was therefore conceivable that hypertonicity might exert its general inhibitory effect on exocytosis through the perturbation of the actin skeleton.
To test this hypothesis, we initially characterized the shrinkage-induced changes in the amount and distribution of filamentous actin in neutrophils with the use of rhodamine phalloidin staining. Exposure of the cells to hypertonicity (500 mosmol/kgH2O) caused a sizeable and persistent increase in F-actin, as verified by flow cytometry (Fig. 2A). The phalloidin fluorescence of shrunken neutrophils increased by >100% within a minute and remained at this elevated level or continued to slightly rise further as long as hyperosmolarity was maintained. This kinetic is markedly different from the FMLP-elicited two-phase response (11), which is characterized by a rapid and even greater initial increase, followed by a gradual return to baseline values (Fig. 2A). When the cells were exposed to both hypertonicity and FMLP, the ensuing rapid rise in F-actin was of similar magnitude as that observed with FMLP alone; however, the depolymerization phase was entirely missing. Thus while FMLP remained capable of inducing actin polymerization even in the presence of hypertonicity, the transient nature of the response was abolished. As a result, after 20 min of stimulation (i.e., when exocytosis was measured), actin content was more than three times higher in cells treated with hypertonicity and FMLP compared with those treated with FMLP alone. We observed similar changes in cytoskeleton-associated actin by Western blotting, confirming that the fluorimetric findings were not due to optical artifacts that might be caused by cell shrinkage (Fig. 2B).
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Relationship between hypertonicity-induced actin polymerization and the inhibition of exocytosis. To assess whether the osmotically induced actin changes and the concomitant block of exocytosis might be causally linked, we applied two approaches: first, we attempted to set differential prestimulus actin levels and measure whether this parameter correlated with the subsequent stimulus-induced exocytosis; and second, we tested whether pharmacological interference with the hypertonic actin response would restore degranulation.
To obtain varying basal actin levels in resting neutrophils, we took advantage of the reversibility of the hyperosmotic actin polymerization.2 To determine the kinetics of the posthypertonic decay of F-actin levels, cells were treated with a 500-mosmol/kgH2O solution for 5 min, followed by rapid return to isotonic medium. Figure 3A shows that after reestablishment of isotonic conditions, the intracellular F-actin level gradually reverted to near baseline values over the course of 5-10 min. At various time points during this actin recovery, the same sample of cells was challenged with FMLP, and the subsequent exocytosis was quantified. To minimize the time necessary to detect exocytosis itself, we followed the expression of CD66b, because 3 min were sufficient to induce a sizeable mobilization of secondary vesicles by FMLP as a stimulus. As shown in Fig. 3A, 3 min after returning to isotonic medium, the total F-actin content was still elevated and only a slight increase in exocytosis was measured. After 5 min in the isotonic treatment, actin depolymerized to near normal levels, and the FMLP-induced exocytosis was gradually resumed. Thus the curves depicting prestimulus F-actin levels and poststimulus exocytosis were almost mirror images of each other, with a slight time delay that occurred in the degranulation response compared with the cytoskeletal change. To better visualize this relationship, Fig. 3B shows the percent change in F-actin content plotted against the percent change in CD66 expression. Clearly, the two phenomena are inversely related in a near hyperbolic manner. Essentially similar observations were made with PMA or A-23187 as stimuli and CD66 or CD35 as exocytosis markers (not shown). Taken together, both the hypertonicity-triggered actin polymerization and the inhibition of exocytosis are fully reversible under isotonic conditions, and their recovery follows a similar time course.
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The effect of pharmacologically induced actin polymerization
on exocytosis under isotonic conditions.
The following experiments were performed to discern whether the
induction of strong actin polymerization is sufficient to inhibit
exocytosis even in an isotonic environment. To address this question,
we used JK, a membrane-permeable sponge toxin that elicits robust and
irreversible actin polymerization (4). Initially, we
verified that 10 µM JK caused monomeric actin to polymerize into
F-actin in neutrophils. Because JK and phalloidin compete for
the same binding sites, the JK-induced F-actin response cannot be
quantified by flow cytometry (4). Therefore, we assayed actin polymerization by measuring cytoskeleton-associated actin with
the use of Western blotting. As expected, exposure to JK under isotonic
conditions resulted in a marked increase in the cytoskeleton-associated
actin, even higher than the rise caused by hypertonicity (Fig.
6A). Moreover, JK was reported
to induce submembranous F-actin deposition (36, 41),
similar to the effect of hypertonicity. Figure 6, B-D,
shows that the drug completely inhibited the FMLP-triggered exocytic
release of markers from all four neutrophil granules. Similar to the
effect of hypertonicity, the JK-induced block of exocytosis was
independent of the stimulus, whether it was FMLP, PMA, A-23187, or LPS.
However, in contrast to hyperosmolarity, JK did not elicit any change
in the neutrophil volume (Fig. 6E) and did not cause general
phosphotyrosine accumulation (Fig. 6F) or the
phosphorylation of osmosensitive kinases such as Syk or p38 (not
shown). Furthermore, JK pretreatment did not affect the FMLP-triggered
tyrosine phosphorylation (Fig. 6F). Thus JK mimicked the
effect of hypertonicity on actin polymerization and on exocytosis
without affecting cell volume or the shrinkage-related signaling. These
data indicate that extensive actin polymerization is sufficient to
prevent exocytosis under isotonic conditions.
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Investigation of the potential involvement of tyrosine
phosphorylation in the hypertonic actin polymerization.
In subsequent experiments, we tested whether the hypertonicity-induced
tyrosine phosphorylation could be an upstream signal for the osmotic
actin polymerization. Osmotic shock results in extensive tyrosine
phosphorylation in neutrophils, including activation of various members
of the Src family, as well as Syk, Pyk2, and p38 (22, 25,
37). To test the potential involvement of these osmosensitive
enzymes in the actin response, we used a variety of kinase inhibitors.
Specifically, we applied the broad-spectrum tyrosine kinase inhibitor
genistein, the Src family inhibitor PP2 that has been shown to prevent
the osmotic activation of Syk and Pyk2 (38), and
SB-203580, a highly specific p38 blocker. As shown on Fig.
7A, none of these compounds
affected the magnitude or the kinetics of the osmotically elicited
increase in F-actin. Moreover, although protein tyrosine
phosphorylation is known to play a crucial role in many FMLP-induced
functions, the FMLP-triggered increase in F-actin remained intact in
the presence of the applied inhibitors (Fig. 7B and Ref.
27). These data suggest that protein tyrosine kinase activation is not
an upstream mediator of the osmotically elicited actin response or a
prerequisite for the FMLP-induced effect.
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The role of ionic strength and cell shrinkage in the hypertonic effect on actin. Actin polymerization is strongly influenced in vitro by concentration of various cations, including K+, Na+, Mg2+, and Ca2+. The increase of ionic strength strongly facilitates actin polymerization in vitro, and this effect is reversed upon reestablishing the original ionic composition. A decrease in cell volume causes a proportional rise in the concentration of the poorly-buffered intracellular ions, such as K+ and Na+. It was conceivable that the elevation in K+ (or Na+) and/or in the total ionic strength of the cytosol could affect actin polymerization. Moreover, in shrunken cells, the concentration of monomeric actin also increases, which may further promote F-actin assembly. Therefore, we designed experiments in which the potential involvement of the major ions, the ionic strength, and cell volume could be separately evaluated.
To establish conditions in which intracellular ionic strength (or species) and cell volume could be manipulated independently, we used KCl-based, intracellular-like media, and nystatin, an ionophore that permeabilizes the membrane selectively to monovalent ions. The basal F-actin levels were not significantly different in cells suspended in isotonic NaCl or KCl medium, and induction of hyperosmolarity by the addition of extra KCl, NaCl, or sucrose was equally effective to induce actin polymerization (Fig. 8A). Because all these osmolytes are poorly permeant, this treatment resulted in equal volume reduction (Fig. 8B) and a parallel rise in intracellular ionic strength. The cells were then incubated in KCl medium supplemented with nystatin. In the presence of the ionophore, the unbalanced intracellular colloid osmotic pressure would drive water entry and consequent cell swelling. To counteract this effect, we added 60 mM sucrose to the medium, which was shown to balance the colloid osmotic pressure, and maintained normal neutrophil volume under these conditions (Fig. 8B). The basal F-actin content of the cells suspended in this medium was somewhat higher than without permeabilization. More importantly, the addition of 100 mM extra KCl caused substantial increase in the F-actin level. This effect was not specific for K+, because addition of NaCl induced a similar response (Fig. 8A). It is important to note that under these conditions, the intracellular ionic strength increased, whereas the cell volume remained constant (Fig. 8B) because the membrane did not pose any barrier against the movement of the added ions. In contrast, the impermeant sucrose, which does not alter ionic strength but causes shrinkage (Fig. 8B and Refs. 25 and 44), failed to elevate the F-actin level. Taking these data together, we conclude that elevated ionic strength (or the total cation and/or Cl
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DISCUSSION |
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The present studies report two novel findings regarding the effect of hypertonicity on neutrophils: 1) hypertonicity induces perturbation of the cytoskeleton that is manifested by increased actin polymerization that occurs predominantly in the submembranous area, and complete abolition of depolymerization that normally follows stimulus-induced F-actin assembly; and 2) actin polymerization is a key contributor to the osmotic inhibition of exocytosis.
The mechanism of hyperosmotic inhibition of exocytosis.
Previous studies by Kazilek et al. (24) have suggested
that the impaired degranulation on hypertonic treatment was primarily due to distortion of the Ca2+ signal. However, these
authors have correctly noticed that this could not be the exclusive
mechanism, because the ionomycin-promoted release of -glucuronidase
(primary granules) was also abrogated, although the
[Ca2+]i-rising effect of the ionophore was
undisturbed. Several additional observations argue against the primary
role of this mechanism. First, in agreement with a previous study by
Junger et al. (22), we also found that moderate
hypertonicity does not abolish the FMLP-induced Ca2+ signal
(not shown), yet it eliminates exocytosis. In addition, shrinkage
itself induces elevation in the resting Ca2+ without
causing exocytosis. Finally, hypertonicity prevents exocytosis triggered by stimuli that do not elicit or require a Ca2+
signal (e.g., PMA). Our systematic investigation has clearly indicated
that osmotic shock interferes with the release of all four granule
types, independent of the stimulus applied. This observation supports
the notion that hypertonicity brings about a general block in the
exocytic process itself that affects step(s) distal to the various
signaling events initiating granule release.
The influence of hypertonic actin polymerization on other functions. The dramatic effect of hypertonicity on the physiological polymerization/depolymerization cycle is expected to influence other functions or signaling events as well. For example, this phenomenon may be responsible for the hypertonic inhibition of locomotion. Pseudopodium formation is believed to require the initial disassembly of the cortical skeleton at a single locale, followed by the appearance of F-actin in the newly formed protrusion (7). Furthermore, rigid cortical actin organization may affect receptor signaling, too: actin polymerization was suggested to separate the FMLP receptors from the G proteins, thereby facilitating the termination of the signal (21). The reported alteration of the Ca2+ signal may also be related to this phenomenon. Alternatively, the Ca2+ release, and/or the store-operated Ca2+ entry pathways, may be directly regulated by the cytoskeleton. In favor of this notion, CB, and recently LB, were found to increase the Ca2+ signal (14). Our unpublished data indicate that LB substantially prolongs the FMLP-triggered Ca2+ rise, suggesting that the influx of external Ca2+ may be specifically sensitive to depolymerization. This finding is in good agreement with the novel concept that a secretion-like, JK-sensitive process mediates the capacitative Ca2+ influx (36).
Actin polymerization (together with p38 activation) (38) may play an important role in the osmotically induced shedding of L-selectin as well. This suggestion is supported by the findings of Middelhoven et al. (33) who reported that JK induces a loss of L-selectin from neutrophils, and by our preliminary observation that LB inhibits the shrinkage-induced L-selectin shedding. The hypertonic actin polymerization appears to be an important common mechanism that accounts for a variety of hypertonic effects including impaired exocytosis, chemotaxis, rolling, adhesion, and transmigration. However, other hypertonic responses, such as the activation of Na+/H+ exchanger or tyrosine phosphorylation, are independent of the actin changes.What is the mechanism of the osmotically triggered actin polymerization? Although this question remains a topic for future studies, two mutually nonexclusive mechanisms can be considered: 1) direct or indirect physicochemical effects, and 2) activation of signaling pathways controlling actin dynamics. Under in vitro conditions, the dehydration of actin per se has been reported to facilitate polymerization (16). Increased ionic strength has a similar effect. During osmotic shrinkage, both of these conditions occur and may contribute to the overall polymerization. Nevertheless, direct physicochemical effects on actin are unlikely to completely account for the in situ polymerization because 1) the ends of the filaments are capped, and the vast majority of monomeric actin is sequestered by specific proteins, and 2) actin polymerization occurs preferentially in membrane-associated areas. It is conceivable that changes in ionic conditions (and in hypertonically induced signaling pathways) induce the dissociation of actin capping proteins such as gelsolin and CapZ (31), whose binding affinity was shown to depend on both KCl and PIP2 concentration, and therefore may be primarily affected at near-membrane areas. Similar mechanisms may alter the major sequestering proteins (profilin, thymosin) as well.
Considering the role of hypertonicity-induced signaling, tyrosine phosphorylation seemed a plausible candidate, because many osmosensitive kinases (e.g., Src family, Syk, Pyk2, and p38) are known to act on the cytoskeleton (47). Moreover, we have recently shown that osmotic stress induces the Src family-dependent phosphorylation of cortactin, a prominent cortical actin cross-linker (23), and hypertonicity has been reported to induce robust tyrosine phosphorylation of actin in Dyctiostelium (48). Nevertheless, activation of the osmosensitive tyrosine kinases proved to be neither sufficient nor necessary for the hypertonicity-induced actin polymerization. Clearly, osmotic shock simultaneously initiates these two parallel responses. We observed that the originally inhibited degranulation was often potentiated after reestablishing isotonicity. Presumably, the facilitating effect of tyrosine phosphorylation was able to manifest after the sterical hindrance was removed. Other alternative signaling pathways include changes in phosphoinositide metabolism and activation of small G proteins. Osmotic stress induces the formation of various phosphoinositides (10), which in turn can induce actin polymerization. Moreover, changes in the membrane structure might expose previously hidden PIP2 molecules. It is noteworthy that hypoosmotic shock induced the rapid dissociation of pleckstrin homology domain-containing proteins from the membrane (15). Small G proteins (e.g., Rac or CDC42) are major regulators of the actin skeleton, and recent studies indicate that they can be translocated to the membrane on shear stress (29) or shrinkage (A. Kapus, unpublished observations). Moreover, some of their downstream targets, such as p21-activated protein kinase, are known to be stimulated by hypertonicity (6). Recent work indicates that small G protein-mediated signaling affects profilin and cofilin, which induce actin polymerization and inhibit depolymerization (9). Future studies should address whether these intriguing mechanisms participate in the hypertonic actin response. ![]() |
ACKNOWLEDGEMENTS |
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The authors are indebted to Dr. S. Grinstein for providing the opportunity to use the Coulter counter and the fluorescence microscope. We thank Hilary Christensen for excellent technical assistance with the electron microscopy studies.
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FOOTNOTES |
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This work was supported by a research fellowship from the Heart and Stroke Scientific Research of Canada (to S. B. Rizoli) and grants from the Medical Research Council of Canada (to A. Kapus and O. D. Rotstein), the Crann Memorial Trust, and the Counnaught Fund of the University of Toronto (to A. Kapus). A. Kapus is a scholar of the Medical Research Council of Canada.
Address for reprint requests and other correspondence: A. Kapus, Toronto General Hospital, Dept. of Surgery, 101 College St., CCRW 2-850, Toronto, Ontario, Canada M5G 1L7 (E-mail: akapus{at}transplantunit.org).
1 Because a part of gelatinase is localized in the secondary granules, exocytosis of these vesicles can contribute to the overall release. However, hypertonicity completely abolished gelatinase release, indicating that the excytosis of the tertiary granules was also inhibited.
2 It is worth noting that by this approach, the impact of the osmotically responsive actin pool can be specifically tested.
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. §1734 solely to indicate this fact.
Received 6 January 2000; accepted in final form 4 April 2000.
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