1Division de Néphrologie, Fondation pour Recherches Médicales, and 2Département de Pathologie, Centre Médical Universitaire, CH-1211 Genève 4, Switzerland
Submitted 7 January 2003 ; accepted in final form 22 April 2003
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ABSTRACT |
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mitogen-activated protein kinase; osmolarity; kidney medulla; cell volume
Hypertonic conditions activate MAPK, which is an important signal transducer linking signals from the cell surface to the nucleus. MAPKs are serine/threonine kinases activated by a cascade of kinases involving two upstream kinases, MAPKKK and MAPKK (48). In mammalian cells, MAPKs are divided into three families, each responding to distinct extracellular stimuli: ERK 1 and 2, JNK (also known as stress-activated protein kinases 1), and p38 kinases (or stress-activated protein kinase 2). In our previous study, we showed that cell shrinkage, rather than intracellular hypertonicity, triggers the activation of ERK and p38 kinase in rat MTALs (35). MAPK activation levels were dependent on the osmolyte used to increase extracellular osmolality. Hyperosmotic NaCl induced cell shrinkage and activated ERK and p38 kinase but not JNK. In comparison, hyperosmotic sucrose induced even greater cell shrinkage and stronger activation of ERK and p38 kinase and also activated JNK but to a lesser extent. By contrast, hyperosmotic urea altered neither cell volume nor MAPK activity. Both hypertonic NaCl and sucrose triggered cellular RVI that restored, almost completely for NaCl and partially for sucrose, the initial cellular volume. Inhibition of p38 kinase decreased the efficiency of RVI, implying a major role of this kinase in this process, whereas inhibition of ERK did not alter RVI.
Modifications of cellular architecture related to hypertonicity-induced cell shrinkage are associated with a reorganization of the architecture of the actin cytoskeleton and with changes in the F-actin-G-actin equilibrium (13, 18, 19, 32, 39). Specific cytoskeleton components may sense cell volume decrease and initiate signaling cascades leading to RVI. In addition, signal transduction cascades leading to remodeling of the actin cytoskeleton and to MAPK activation share some common elements. For instance, small G proteins of the Rho family, such as Cdc42 and Rac 1, are involved in both actin cytoskeleton remodeling through filipodia and lamellipodia formation (45) and in signaling events leading to p38 kinase activation (1, 51). Activation of p38 kinase may, in turn, control actin cytoskeleton dynamics through the activation of downstream kinases such as MAPKAP kinase 2/3 or PRAK, which phosphorylate HSP 25/27 (17, 28, 37), a small heat shock protein that modulates actin polymerization (27). The actin cytoskeleton may also control the activity of ion transporters, leading to intracellular NaCl uptake and secondary water influx, either directly, through F-actin/G-actin ratio dynamics (9, 10), or indirectly, through binding of signaling modules (50) and/or modulation of endocytotic-exocytotic events (42). This study was therefore undertaken to investigate the relationship between actin cytoskeleton remodeling and RVI in rat MTAL.
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MATERIALS AND METHODS |
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Preparation of MTAL suspensions. The two kidneys were perfused
with ice-cold incubation solution without collagenase. The inner stripes of
the outer medulla were excised, minced on ice, and fragments of medullary
tubules were obtained by gentle pressure through nylon filters with a pore
size decreasing from 150 to 100 µm. After centrifugation, the pellet was
resuspended in ice-cold oxygenated (95% O2-5% CO2)
incubation solution. As controlled under a stereomicroscope, MTALs account for
90% of the tubule fragments in this preparation. Therefore, it will be
referred as MTAL suspension.
Determination of the Triton X-100-soluble and -insoluble actin
fractions. Estimation of actin polymerization level was performed by
determining the Triton X (TX)-100-soluble/TX-100-insoluble actin ratio.
Indeed, it is largely admitted that F-actin, i.e., polymerized actin, is
contained in the TX-100-insoluble fraction and that G-actin, i.e., monomeric
actin, is contained in the TX-100-soluble fraction
(16). After 1-h preincubation
at 30°C in isotonic incubation solution with or without addition of drugs,
MTAL suspensions were incubated at 37°C for 1 to 30 min under isosmotic or
hyperosmotic (addition of 300 mosM/l NaCl, sucrose, or urea) conditions.
Incubation was stopped by cooling and centrifugation at 6,000 g for 5
min at 4°C. The pellet was saved and 20 µl of ice-cold lysis buffer (20
mM Tris · HCl, 2 mM EGTA, 2 mM EDTA, 30 mM NaF, 30 mM
Na4O7P2, 2 mM Na3VO4, 1
mM AEBSF, 10 µg/ml leupeptin, 4 µg/ml aprotinin, 1% Triton X-100, pH
7.45) were added. After 5 min of centrifugation at 12,000 g, the
supernatant was saved, the pellet was then mixed with fresh lysis buffer, and
after a centrifugation step at 12,000 g, the second supernatant was
saved. The final pellet was then suspended in sample buffer and an equal
volume of sample buffer was added to the pooled supernatants. The proteins
from pooled supernatants and pellet were then separated by 10% SDS-PAGE and
transferred to a polyvinylidene difluoride (PVDF) membrane (Immobilon-P,
Millipore, Waters, MA), and -actin was detected by immunoblot using a
monoclonal anti-
-actin antibody (AC-15, Sigma, St. Louis, MO) at
1:40,000 dilution (vol/vol). After incubation with anti-mouse IgG coupled to
horseradish peroxidase (Transduction Laboratories, Lexington, UK) at 1:10,000
dilution (vol/vol), immunoreactivity was detected by chemiluminescence using
the Super Signal Substrate method (Pierce, Rockford, IL). Results were
quantified under conditions of linearity by integration of the density of
total area of each band using a video densitometer and Image-Quant software
(Molecular Dynamics, Sunnyvale, CA). Results are expressed as a percentage of
the control optical density (isotonic medium) and are means ± SE.
Determination of the phosphorylation level of ERK and p38 kinase. After 1-h preincubation at 30°C in isotonic incubation solution with or without addition of drugs, MTAL suspensions were incubated at 37°C for 10 min under isosmotic or hyperosmotic (addition of 300 mosM/l NaCl) conditions. Incubation was stopped by cooling and centrifugation at 6,000 g for 5 min at 4°C. After addition of lysis buffer, protein content was measured by the BCA protein assay (Pierce). Equal amounts of protein (50 µg) were separated by 10% SDS-PAGE and transferred to a PVDF membrane (Immobilion-P, Millipore). Phosphorylated ERK and p38 kinase were detected using anti-ERK-P and anti-p38-P kinase rabbit polyclonal antibodies (New England Biolabs, Beverly, MA) at 1:10,000 dilution (vol/vol). After incubation with anti-rabbit IgG coupled to horseradish peroxidase (Transduction Laboratories) at 1:10,000 dilution (vol/vol), immunoreactivity was detected by chemiluminescence, and results were quantified and expressed as described above.
Determination of MTAL cellular volume. A pool of three isolated
MTALs was transferred into the concavity of a bacteriological slide coated
with dried BSA. After 1-h preincubation at 30°C in isosmotic incubation
solution with or without addition of drugs, MTALs were incubated at 37°C
for 1 to 30 min under isosmotic or hyperosmotic (addition of 300 mosM/l NaCl)
conditions. After preincubation at 30°C in isosmotic incubation solution,
tubules were incubated in isosmotic or hyperosmotic incubation solutions with
or without drugs. MTALs were visualized with an inverted microscope, and
photographs of the same tubules were taken at the end of the preincubation
period and after incubation. MTAL volume (V) was calculated from the measured
radius (R) and length (L) of the tubules at a 1,000-fold
magnification using the formula V = R2 x
L. Because the lumen is collapsed in nonperfused tubules, we assumed
that MTAL volume measurement is an appropriate estimate of MTAL cellular
volume. Results are expressed as a percentage of the control volume (end of
the preincubation period) and are means ± SE.
Fluorescence microscopy. MTAL suspensions were preincubated at 30°C for 1 h with or without drugs and then incubated at 37°C under isosmotic or hyperosmotic (addition of 300 mosM/l NaCl or sucrose) conditions. Tubules were then cytocentrifuged on glass slides using a cytospin (70 g, 5 min in incubation solution supplemented with 1% BSA) and fixed with 3.7% paraformaldehyde for 10 min at room temperature. After three washes in PBS, fixed tubules were permeabilized with 0.1% Triton X-100 for 1 min at room temperature. After a new series of three washes in PBS, specimens were incubated with phalloidin Alexa-488 (dilution: 1:100 in PBS; Molecular Probes, Eugene, OR) for 1 h at room temperature. Specimens were observed with a Zeiss Axiophot microscope (Carl Zeiss, Jena, Germany) equipped with an oil-immersion plan-neofluar x40:1.3 objective. Images were acquired with a high-sensitivity, high-resolution color camera (Axiocam, Carl Zeiss). Pictures were printed with a digital pictrography 4000 printer (Fujifilm, Tokyo, Japan).
Statistical analysis. Statistical analysis of variations of TX-100-insoluble/TX-100-soluble actin and cellular volume was done by ANOVA. Statistical analysis of variations of anti-P-ERK and P-p38 kinase immunoreactivity was done using the Kruskall-Wallis test. P values <0.05 were considered significant.
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RESULTS |
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Figure 1 shows that increasing extracellular osmolarity up to 600 mosM/l by addition of NaCl, sucrose, or urea rapidly increased the proportion of TX-100-insoluble actin, i.e., F-actin, with a peak observed after 1- to 3-min incubation at 37°C. The TX-100-insoluble/TX-100-soluble actin ratio then returned close to its basal level after 10-min incubation. These rapid variations in cellular TX-100-insoluble actin content were followed by a progressive increase in proportion of TX-100-insoluble actin above the basal levels in samples incubated up to 30 min in the presence of hyperosmotic NaCl (TX-100-insoluble/TX-100-soluble actin; isosmotic: 1.46 ± 0.18; NaCl: 2.46 ± 0.30; P < 0.05). This increase in proportion of TX-100-insoluble actin was sustained for at least 60 min (data not shown). The progressive increase in proportion of TX-100-insoluble actin was more pronounced after 30 min in the presence of hyperosmotic sucrose (TX-100-insoluble/TX-100-soluble actin; isotonic: 2.10 ± 0.37; sucrose: 5.98 ± 1.03; P < 0.05) compared with hyperosmotic NaCl (Fig. 1, A and B). In contrast, for incubation periods ranging from 10 to 30 min, hyperosmotic urea did not induce significant variations in the TX-100-insoluble/TX-100-soluble actin ratio (isotonic 30 min: 1.75 ± 0.11; urea 30 min: 1.60 ± 0.12; not significant; Fig. 1C). As shown previously (35), both hyperosmotic NaCl and sucrose rapidly induced cell shrinkage with a maximal decrease in cellular volume observed after 10 min of incubation (% of initial cellular volume; NaCl: 68.98 ± 2.21; sucrose: 65.47 ± 0.58). After 30-min incubation in the presence of hyperosmotic NaCl or sucrose, a partial recovery of the initial cellular volume was observed (NaCl: 89.14 ± 2.24; sucrose: 81.81 ± 5.94; Fig. 1, A and B). In contrast, hyperosmotic urea did not significantly alter cellular volume (Fig. 1C). These results show that, in MTAL cells, acute extracellular hyperosmolality induces polyphasic actin cytoskeleton remodeling reflected by the observed changes in the F-actin/G-actin ratio. However, sustained actin polymerization reflected by the progressive increase in cellular F-actin content was only observed in response to osmolytes inducing cell shrinkage and this event occurred concomitantly with the partial recovery of the initial cellular volume.
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The effect of extracellular hyperosmolality on actin cytoskeleton organization was assessed by fluorescence microscopy on isolated rat MTALs incubated at 37°C for 30 min under isosmotic or hyperosmotic conditions. As shown by Fig. 2A, rat MTAL cells incubated under isosmotic conditions exhibited a dense cortical F-actin ring delineating the cell periphery and a sparse diffuse network of F-actin bundles. After exposure of MTALs to hyperosmotic NaCl, the cortical F-actin ring was thinner and the diffuse F-actin network was more developed compared with tubules incubated under isosmotic conditions (Fig. 2B). This redistribution of F-actin was more pronounced after incubation of tubules in the presence of hyperosmotic sucrose (Fig. 2C). In contrast, hyperosmotic urea did not induce any significant change in F-actin distribution (Fig. 2D). Therefore, the sustained actin polymerization phase observed in response to hyperosmotic NaCl and sucrose was associated with a redistribution of F-actin from the cortical F-actin ring to a diffuse network of F-actin bundles. In contrast, the early actin polymerization phase was not associated with apparent F-actin redistribution (data not shown).
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Interfering with actin polymerization and remodeling altered cellular volume recovery under hypertonic conditions. The actin cytoskeleton is a highly dynamic structure that undergoes constant remodeling, consisting of spatially and temporally regulated polymerization and depolymerization of preexisting filaments as well as nucleation and branching of new actin filaments. New pharmacological tools derived from marine sponges (40) were used to study the role of actin polymerization-depolymerization and generation of new actin filaments on cellular volume recovery under hypertonic conditions. Jasplakinolide binds to both ends of actin filaments preventing their depolymerization and also causes rapid nucleation of actin polymerization (5, 40). As expected from its mechanism of action, 10 µM jasplakinolide (Calbiochem, San Diego, CA) increased the proportion of TX-100-insoluble actin in tubules incubated under hypertonic conditions for 30 min (TX-100-insoluble/TX-100-soluble actin; NaCl: 2.87 ± 0.49; NaCl + jasplakinolide: 4.02 ± 0.45; P < 0.05; Fig. 3A). In addition, fluorescence microscopy revealed that jasplakinolide-treated tubules exhibited a very dense F-actin network with diffuse thick and short actin-rich structures, i.e., actin clumps, throughout the cytoplasm (Fig. 3B). Therefore, jasplakinolide efficiently increases the actin polymerization in rat MTAL cells. Swinholide A inhibits actin filament nucleation and elongation (6, 40), thereby preventing stimulus-induced actin polymerization without affecting the intact actin network (46). Measurement of the partition of actin between TX-100-soluble and -insoluble fractions showed that, in agreement with its pharmacological properties, 50 µM swinholide A (Calbiochem) moderately decreased the amounts of TX-100-insoluble actin measured after 30-min incubation under hypertonic conditions (TX-100-insoluble/TX-100-soluble actin; NaCl: 2.87 ± 0.49; NaCl + swinholide: 2.17 ± 0.28; not significant; Fig. 3A). Fluorescence microscopy, however, showed that reorganization of the actin cytoskeleton induced by hyperosmotic NaCl was largely prevented by swinholide A (compare a and c, Fig. 3B). Indeed, most F-actin remained in the cortical ring and the density of the diffuse F-actin network was unchanged compared with MTALs incubated under isotonic conditions. Therefore, swinholide A does not significantly alter actin cytoskeleton organization but prevents its hypertonicity-induced remodeling in rat MTAL cells. Latrunculin B sequesters monomeric actin and decreases G-actin availability, resulting in actin filament depolymerization (40, 41). Consistent with its actin-depolymerizing properties, 100 µg/ml latrunculin B (Calbiochem) induced a large decrease in proportion to TX-100-insoluble actin with respect to control after 30-min incubation under hypertonic conditions (TX-100-insoluble/TX-100-soluble actin; NaCl: 2.87 ± 0.49; NaCl + latrunculin: 0.33 ± 0.09; P < 0.01; Fig. 3A). In addition, fluorescence microscopy revealed that latrunculin B disorganized the actin cytoskeleton. The cortical F-actin ring became irregular and discontinuous, and the diffuse F-actin network was almost completely disrupted (compare a and d, Fig. 3B). Therefore, latrunculin B potently depolymerizes the actin cytoskeleton in rat MTAL cells.
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The role of actin polymerization or depolymerization and of the generation of new actin filaments of cellular volume recovery was assessed using actin-interfering drugs. Figure 4A shows that jasplakinolide, which induces actin polymerization and prevents actin depolymerization, almost completely prevented the partial recovery of the initial cellular volume observed after incubation of isolated MTALs for 30 min under hypertonic conditions but in the absence of drug (% of initial volume; NaCl: 88.48 ± 4.71; NaCl + jasplakinolide: 74.65 ± 0.71; P < 0.05). Swinholide A (Fig. 4B), which prevents the generation of new actin filaments, increased cell shrinkage after 5-min incubation under hypertonic conditions (% of initial volume; NaCl: 69.61 ± 3.90; NaCl + swinholide: 51.85 ± 4.86; P < 0.05) and decreased the extent of recovery of the initial cellular volume observed after 30 min in the presence of hyperosmotic NaCl (% of initial volume; NaCl: 88.48 ± 4.71; NaCl + swinholide: 75.90 ± 3.09; P < 0.05; Fig. 4B). Finally, latrunculin B, which depolymerizes the actin cytoskeleton, largely attenuated cell shrinkage in response to 5-min incubation with hyperosmotic NaCl (% of initial volume; NaCl: 69.61 ± 3.90; NaCl + latrunculin: 80.84 ± 4.28; P < 0.05) and allowed a full recovery of cellular volume after 30-min incubation (% of initial volume; NaCl: 88.48 ± 4.71; NaCl + latrunculin: 102.54 ± 4.94; P < 0.05; Fig. 4C). Thus, in rat MTALs, inhibition of actin depolymerization and generation of new actin filaments decrease the efficacy of RVI, whereas actin depolymerization potentiates cell volume recovery after an acute hypertonic challenge.
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Interfering with actin polymerization and remodeling did not alter MAPK activation by hyperosmotic NaCl. The role of actin cytoskeleton remodeling in MAPKs activation in response to extracellular hypertonicty was assessed by measurement of the phosphorylation level of ERK and p38 kinase in the absence or presence of actin-interfering drugs. Figure 5 shows that the increases in phosphorylation levels of ERK and p38 kinase observed after incubation at 37°C for 10 min under hypertonic conditions in the presence of jasplakinolide, or swinholide A or latrunculin B, were similar to those induced by hyperosmotic NaCl alone. Similarly, ERK and p38 kinase phosphorylation levels were not altered by actin-interfering drugs in MTALs incubated under isotonic conditions (data not shown). Therefore, activation of ERK and p38 kinase in response to extracellular hypertonicity is independent of actin cytoskeleton remodeling.
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p38 Kinase was involved in cellular volume recovery and actin cytoskeleton reorganization following extracellular hypertonic challenge. The following experiments were designed to study the role of MAPKs in cellular volume variations and actin cytoskeleton reorganization induced by extracellular hypertonicity. Inhibition of the ERK signaling pathway by 4.104 M PD-98059 (Calbiochem) modified neither cellular volume variation nor TX-100-insoluble/soluble actin ratio profiles in response to hyperosmotic NaCl (data not shown). Inhibition of p38 kinase by 105 M SB-203580 (Calbiochem) slightly increased the maximal extent of hypertonicity-induced cell shrinkage observed after 10 min (% of initial volume; NaCl: 77.14 ± 2.82; NaCl + SB: 65.51 ± 2.21 ± 4.94; P < 0.05) and decreased the efficacy of cell volume recovery after 30 min (% of initial volume; NaCl: 91.28 ± 1.21; NaCl + SB: 78.05 ± 3.11; P < 0.05; Fig. 6A). In addition, SB-203580 attenuated the early increase in proportion to TX-100-insoluble actin observed after 2 min in the presence of hyperosmotic NaCl (TX-100-insoluble/TX-100-soluble actin; NaCl: 2.57 ± 0.43; NaCl + SB: 1.79 ± 0.25; P < 0.05) and abolished the sustained increase in amounts of TX-100-insoluble actin observed after 30-min incubation with hyperosmotic NaCl (TX-100-insoluble/TX-100-soluble actin; NaCl: 2.46 ± 0.30; NaCl + SB: 1.50 ± 0.09; P < 0.01; Fig. 6B). Similar results were obtained in the presence of hyperosmotic sucrose (data not shown). As previously shown (35), SB-203580 did not alter MTAL cellular volume measured under isotonic conditions. Thus both cellular volume recovery and sustained actin polymerization phase, which occur concomitantly, are dependent on p38 kinase activity.
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Inhibition of p38 kinase activity and swinholide A both prevent F-actin redistribution in response to extracellular hypertonicity. SB-203580, which inhibits p38 kinase, and swinholide A, which prevents generation of new actin filaments, both inhibited the sustained actin polymerization and decreased the efficacy of RVI in response to extracellular hypertonicity. We therefore assessed by fluorescence microscopy the effect of SB-203580 and swinholide A on F-actin redistribution following hypertonic challenge. As shown by Fig. 7, SB-203580 and swinholide A strongly attenuated the hypertonicity-induced redistribution of F-actin from the dense cortical F-actin ring to the diffuse network of F-actin bundles (compare Fig. 7, A and B), compared with tubules incubated in the presence of hyperosmotic NaCl alone (compare Fig. 7, B-D). These results suggest that SB-203580 and swinholide A share the same mechanism of inhibition of actin cytoskeleton remodeling.
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DISCUSSION |
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Rat MTAL cells, which are physiologically exposed to large variations in interstitial osmolality (23), exhibit a polyphasic actin polymerization profile in response to extracellular hyperosmolality (see Fig. 1). The initial rapid actin polymerization and depolymerization phases were not associated with detectable actin filament redistribution and were shared by hyperosmotic challenges induced by NaCl, sucrose, and urea. Because urea altered neither cellular volume nor MAPK activity (35), the transient phases of actin polymerization-depolymerization are obviously independent of cell shrinkage and MAPK activation. In whole organisms, the interstitial osmolality of the kidney medulla increases progressively under antidiuresis conditions, whereas under ex vivo experimental conditions used in this study, extracellular osmolality increased abruptly. We therefore cannot exclude the possibility that the observed rapid changes in the levels of actin polymerization observed during the first 10 min of incubation are due to an acute increase in intracellular osmolality. Exposure of rat MTAL cells to hyperosmotic NaCl and sucrose, but not urea, induced a progressive actin polymerization phase (from 10- to 30-min incubation) and redistribution of F-actin from a dense cortical ring to a diffuse network of F-actin bundles that may rely on cell shrinkage and subsequent MAPK activation (35). Actin polymerization and redistribution of F-actin were more pronounced in response to hyperosmotic sucrose, which decreases cellular volume and activates MAPKs to a larger extent than NaCl. In contrast, the sustained phase of actin polymerization and the redistribution of F-actin were not observed in the presence of hyperosmotic urea, which does not alter cell volume and MAPK activity. The abolition of sustained actin polymerization and redistribution of F-actin by a specific p38 kinase inhibitor further support this interpretation (see Figs. 6 and 7).
Our results suggest that actin cytoskeleton remodeling is dependent on p38 kinase activation (see Figs. 6 and 7). The p38 kinase-dependent actin cytoskeleton remodeling may be mediated, at least in part, through phosphorylation of HSP25/27, a small heat shock protein that modulates actin polymerization (27). Phosphorylated HSP25/27 promotes actin polymerization, whereas its nonphosphorylated form is inhibitory (2, 8, 25, 31, 38). In intact cells, activated p38 kinase phosphorylates and activates MAPKAP kinase 2/3, which in turn phosphorylates HSP25/27 (17, 28, 37) and thereby promotes redistribution of HSP25/27 from the cytoplasm to the actin cytoskeleton (52). However, in addition to the MAPK pathway, cell shrinkage increases tyrosine phosphorylation of a subset of proteins including nonreceptor tyrosine kinases and cytoskeleton-associated proteins (20, 21, 24). Therefore, the tyrosine kinase pathway may also participate in the actin cytoskeleton remodeling induced by extracellular hypertonicity.
In addition to native MTAL cells, remodeling of the actin cytoskeleton in response to extracellular hyperosmolality has been observed in yeast (12), Dictyostelium (53), and cultured mammalian nonepithelial cells (13, 18, 19, 32) as well as in epithelial Madin-Darby canine kidney cells (39). Results of the present study and from the literature indicate that actin cytoskeleton remodeling exhibits some degree of cell specificity. In native rat MTAL cells (see Fig. 2) and glial cells (32), hypertonicity induced redistribution of F-actin from the cortical ring to a diffuse network of actin bundles, whereas in fibroblasts and HL60 cells, a densification of the peripheral actin ring was observed (13, 18). Moreover, the sustained actin polymerization phase observed in native rat MTAL cells was absent in cultured HL60 cells (19). These different patterns of actin cytoskeleton remodeling are associated with differences in RVI efficacy. Indeed, in contrast to the majority of cells exhibiting little or no RVI, MTAL cells undergo robust RVI (35, 43).
The temporal relationship between RVI and actin cytoskeleton reorganization, taken together with results obtained with actin-interfering drugs, suggests that both cortical F-actin depolymerization and de novo actin polymerization resulting in the generation of a diffuse network of F-actin bundles play an important role in the RVI of MTAL cells. Results of the present study indicate that whole cell actin depolymerization with latrunculin B facilitates RVI, whereas global inhibition of actin depolymerization by jasplakinolide antagonizes RVI (see Figs. 3 and 4). These results suggest that depolymerization of F-actin is required for RVI in MTAL epithelial cells. In addition, fluorescence microscopy imaging shows that RVI is associated with reduced cortical F-actin staining (see Fig. 2), suggesting that the F-actin depolymerization process involved in RVI specifically takes place at the level of the cortical F-actin ring in MTAL epithelial cells. This result contrasts with those obtained in nonepithelial cells (HL60) that exhibit a densification of the cortical F-actin ring in response to hypertonicity but which do not undergo RVI (13, 18). On the other hand, our results show that swinholide A or SB-203580, an inhibitor of p38 kinase, prevented the hypertonicity-induced generation of diffuse F-actin bundles and reduced the efficacy of RVI (see Figs. 4 and 6). These results suggest that, in addition to cortical F-actin ring depolymerization, the generation of a dense diffuse network of F-actin bundles facilitates RVI. At fist glance, this finding contrasts with the effect of jasplakinolide, which increases the actin polymerization level and prevents the RVI. It should be noticed, however, that jasplakinolide also increased actin polymerization at the level of the cortical F-actin ring, an effect that most likely antagonizes RVI.
It is well established that RVI is associated with ion transporter activation including Na-K-2Cl cotransporter, Na/H exchanger, Cl/HCO3 exchanger, and Na-K-ATPase (4, 15, 29, 43, 47), which might, at least in part, be dependent on cortical actin polymerization level. For instance, inhibition of F-actin depolymerization by phalloidin or jasplakinolide impairs the activation of the Na-K-2Cl cotransporter by cAMP in MTAL cells (49). Conversely, depolymerization of F-actin by cytochalasin D stimulates Na-K-2Cl cotransporter in intestinal cells (30). Actin polymerization may control the activity of ion transporters in different ways. A shift in F-actin-G-actin equilibrium toward G-actin may stimulate the activity of specific ion transporters, as shown for Na-K-ATPase (9) and epithelial Na channels (3, 10). On the other hand, depolymerization of the cortical F-actin ring may promote the exocytosis of ion, solute, and water transporters as demonstrated for the Na/H exchanger NHE3 (11), volume-sensitive Cl channels (33), the glucose transporter GLUT4 (26), and the water channel aquaporin-2 (22). The results of the present study indicate that, at the level of the whole cell, the equilibrium between actin polymerization and depolymerization is shifted toward actin polymerization during RVI (see Fig. 1). This polymerization process results in the generation of a dense and diffuse network of F-actin bundles (see Fig. 2) that may play a functional role in the defense against cell shrinkage. Indeed, inhibition of the sustained actin polymerization phase and the densification of the diffuse F-actin network by swinholide A and SB-203580 both increased the extent of maximal cell shrinkage and reduced the efficacy of RVI in response to hypertonicity (see Figs. 4, 6, and 7). This effect might be partly achieved through mechanical constraints exerted on the cell membrane, as described for lamellipodia or filipodia formation (44). It occurs, however, most likely indirectly via spacial control of signaling events and/or facilitation of the delivery of ion transporters from intracellular stores to the plasma membrane, as shown for the GLUT4 glucose transporter in response to insulin (46).
Because activation of MAPKs is mediated by cell shrinkage in rat MTAL cells (35), the hypothesis that the actin cytoskeleton may be part of the osmosensing machinery was considered. Our results, however, suggest that actin cytoskeleton remodeling and integrity are not essential for the activation of MAPKs in response to increased extracellular osmolality. Indeed, interfering with neither the polymerization level of actin nor with the generation of new actin filaments decreased the extent of ERK and p38 kinase activation in response to extracellular hypertonicity (see Fig. 5). Therefore, alternative mechanisms such as an increase in cytoplasmic concentration of macromolecules (34) or aggregation of growth factor and cytokine receptors leading to their ligand-independent activation (36) have to be considered.
In conclusion, we showed that an acute extracellular hypertonic challenge induces actin cytoskeleton remodeling consisting of F-actin redistribution from a cortical ring to a diffuse network of F-actin bundles in native rat MTAL cells. We propose the following working hypothesis summarized by Fig. 8. Cell shrinkage induces p38 kinase activation, which in turn, promotes cortical F-actin ring depolymerization and generation of a dense diffuse network of F-actin bundles that both promote RVI most likely through modulation of ion transporter activity. Further investigation is required to identify the molecular players involved in actin cytoskeleton remodeling. In addition, the role of actin cytoskeleton remodeling in the control of the activity and/or abundance of ion transporters involved in RVI remains to be determined.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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REFERENCES |
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