1 Laboratory of Physiology, Katholieke Universiteit Leuven, Campus Gasthuisberg, B-3000 Leuven, Belgium; 2 Institute of Experimental and Clinical Pharmacology and Toxicology, 79104 Freiburg, Germany; 3 Department of Medical Physiology, The Panum Institute, University of Copenhagen, 2200 Copenhagen; and 4 Biochemical Department, August Krogh Institute, University of Copenhagen, 2100 Copenhagen, Denmark
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ABSTRACT |
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Cell swelling triggers in most cell types
an outwardly rectifying anion current,
ICl,swell, via volume-regulated anion channels (VRACs). We have previously demonstrated in calf pulmonary artery endothelial (CPAE) cells that inhibition of the Rho/Rho kinase/myosin light chain phosphorylation pathway reduces the swelling-dependent activation of ICl,swell. However, these
experiments did not allow us to discriminate between a direct activator
role or a permissive effect. We now show that the Rho pathway did not
affect VRAC activity if this pathway was activated by transfecting CPAE
cells with constitutively active isoforms of G (a Rho activating
heterotrimeric G protein subunit), Rho, or Rho kinase. Furthermore,
biochemical and morphological analysis failed to demonstrate activation
of the Rho pathway during hypotonic cell swelling. Finally,
manipulating the Rho pathway with either guanosine
5'-O-(3-thiotriphosphate) or C3 exoenzyme had no effect on
VRACs in caveolin-1-expressing Caco-2 cells. We conclude that the Rho
pathway exerts a permissive effect on VRACs in CPAE cells, i.e.,
swelling-induced opening of VRACs requires a functional Rho pathway,
but not an activation of the Rho pathway.
cytoskeleton; actin; myosin phosphorylation; RhoA pathway
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INTRODUCTION |
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HYPOTONIC CELL SWELLING
ACTIVATES an outwardly rectifying Cl current
(ICl,swell) via volume-regulated anion channels
(VRACs), which is expressed in nearly every mammalian cell type. The
biophysical features of VRACs are an Eisenman type I permeability
sequence (I
> Br
> Cl
> F
), outward rectification, slow
inactivation at positive potentials, and a single channel conductance
of ~40 pS at positive potentials (for reviews, see Refs.
24 and 34). Because VRACs are also permeable to organic
osmolytes, their activation during cell swelling results in an efflux
of both inorganic and organic osmolytes (13). This
provokes an osmotic efflux of water and restoration of cell volume. In
addition, VRACs have been implicated in other cellular functions such
as mechanotransduction, cell proliferation, and apoptosis
(3, 20, 32).
An intriguing, but largely unresolved, property of VRACs is their
activation mechanism. Under experimental conditions, VRACs are
typically activated by cell swelling evoked either by extracellular perfusion with a hypotonic solution or by intracellular dialysis with a
hypertonic pipette solution. However, several observations indicate
that VRACs can also be activated in the absence of cell swelling, e.g.,
by reducing intracellular ionic strength (7, 40), by
mechanical stimulation (3), or by intracellular
application of guanosine 5'-O-(3-thiotriphosphate) (GTPS)
(41). The isovolumic activation has stirred a controversy
about the causal relation between cell volume and VRAC activation. Some
groups have proposed that VRAC activity is primarily controlled by
changes in cell volume but that the cellular volume sensor can be
(de)sensitized by other parameters such as intracellular ionic strength
(7, 8). Alternatively, it has been proposed that a
decrease in intracellular ionic strength (e.g., due to water influx
during hypotonic cell swelling) constitutes the critical trigger for channel activation (40). A strong argument in favor of the
ionic strength mechanism is that single VRACs could be directly
activated by decreased ionic strength (28).
Irrespective of the outcome of this controversy, it has become clear
that activation of VRACs during hypotonic cell swelling depends on the
specific structural and/or molecular architecture of the cell. For
example, hypotonic activation of VRACs is largely deficient in
caveolin-1-deficient Caco-2 cells (a colon carcinoma cell line), but
transient expression of caveolin-1 in these cells recovers VRAC
activity (37). Furthermore, efficient activation of VRACs
requires, at least in some cell types, a functional F-actin cytoskeleton (19, 31). Finally, the transient activation
of VRACs by intracellular application of GTPS points to a modulatory role for GTP-binding proteins (41). Incubation with
Clostridium limosum or C. botulinum C3 toxins
that specifically inactivate Rho GTPases (A/B/C isoforms)
(15) strongly reduced the activation of VRACs by cell
swelling or by intracellular GTP
S (26, 35). Similarly,
C. difficile toxin B, which is another Rho-inhibiting bacterial toxin, inhibits ICl,swell in N1E115
neuroblastoma cells (10). Taken together, these studies
identify Rho GTPases as modulators for VRAC activity.
Rho GTPases (A/B/C isoforms) are GTP-regulated molecular switches that
control contractile activity in smooth muscle and the formation of
stress fibers in noncontractile cell types (33). The Rho
family of GTPases plays important roles in F-actin organization (6). Activation of Rho requires dissociation from a
cytosolic binding partner RhoGDI (guanosine nucleotide dissociation
inhibitor) and exchange of GDP for GTP, which is catalyzed by Rho
guanosine nucleotide exchange factors (RhoGEFs) (6). It
has recently been shown that stimulation of some G protein-coupled
receptors (e.g., thrombin receptor) increases Rho activity (9,
39). The molecular cascade between G protein-coupled receptor
and Rho has also been elucidated: ligand-bound receptors activate
heterotrimeric G12 and/or G13 proteins, of
which the dissociated -subunits (G
12 or
G
13) stimulate RhoGEF (12, 18, 22). Once
activated, Rho exerts its effects via multiple downstream effectors
(6). The Rho-induced stress fiber formation is mediated by
mDia1, which promotes actin polymerization and by Rho kinase, a
serine/threonine protein kinase that induces myosin light chain
phosphorylation by inhibiting myosin light chain phosphatase (6,
33, 42).
We have previously shown that activation of VRACs in calf pulmonary
artery endothelial (CPAE) cells (a macrovascular endothelial cell line)
requires a functional Rho/Rho kinase/myosin light chain phosphorylation
pathway. Indeed, inhibition of Rho, Rho kinase, or myosin light chain
kinase precludes swelling-induced activation of VRAC, whereas
inhibition of myosin light chain phosphatase exerts a potentiating
effect on VRAC (25, 26). In this study, we wanted to
establish whether there is a direct and causal link between
hypotonicity-induced cell swelling, activation of the Rho pathway, and
opening of VRAC. Three implications of this hypothesis were tested:
1) Does activation of the Rho pathway result in an activation of VRAC? 2) Does hypotonic cell swelling activate
the Rho pathway? 3) Is a functional Rho pathway a universal
requirement for VRAC activation? Our data indicate that cell swelling
does not activate the Rho pathway in CPAE cells and, vice versa, that activation of the Rho pathway does not trigger VRAC activation. Furthermore, VRACs are insensitive to GTPS or Rho inhibition by C3
toxins in caveolin-1-expressing Caco-2 cells. We therefore propose that
the Rho pathway exerts a permissive role with respect to VRAC
activation in some cell types such as CPAE cells.
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MATERIALS AND METHODS |
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Cells. We used CPAE (American Type Culture Collection CCL-209) and human colorectal carcinoma cells Caco-2 (ECACC 86010202). The cells were grown in DMEM (Life Technologies, GIBCO) containing 20% FCS, 2 mM L-glutamine, 100 µg/ml streptomycin, and 100 U/ml penicillin, maintained at 37°C in a fully humidified atmosphere of 10% CO2 in air. Passaging of the cells was performed by brief exposure to 0.5 g/l trypsin in a Ca2+- and Mg2+-free solution. Only nonconfluent cells were used in the patch-clamp experiments.
Twenty-four hours after the cells were seeded, they were transiently transfected with the bicistronic pCINeo/IRES-GFP vector (38), containing the cDNA encoding the constitutively active forms of GSolutions. At the beginning of the patch-clamp recordings, a modified Krebs solution containing (in mM) 150 NaCl, 6 KCl, 1 MgCl2, 10 glucose, and 10 HEPES (pH 7.4) with NaOH was replaced by an isotonic Cs+ solution, containing (in mM) 105 NaCl, 6 CsCl, 1 MgCl2, 1.5 CaCl2, 10 glucose, 10 HEPES, and 90 mannitol, adjusted to pH 7.4 with NaOH. The osmolality of the solutions, as measured with a vapor pressure osmometer (Wescor 5500; Schlag, Gladbach, Germany) was 320 ± 5 mosmol/kgH2O. The 25% hypotonic solution (HTS) was obtained by omitting 90 mM mannitol from this Cs+ solution. Pipette solutions contained (in mM) 40 CsCl, 100 cesium aspartate, 1 MgCl2, 1.93 CaCl2, 5 EGTA, 4 Na2ATP, and 10 HEPES, adjusted to pH 7.2 with CsOH (290 mosmol/kgH2O). The concentration of free Ca2+ in this solution was buffered at 100 nM.
In some experiments, 100 µM GTPElectrophysiological recordings.
Transfected green fluorescent cells were visualized in a patch-clamp
setup as described previously (38). Currents were
monitored with an EPC-7 patch-clamp amplifier (List Electronic,
Lambrecht/Pfalz, Germany). Patch electrodes had direct current
resistances between 2 and 6 M. An Ag-AgCl wire was used as reference electrode.
Affinity precipitation of cellular Rho-GTP.
BL21 Escherichia coli bacteria were transformed with the
pGEX3X vector containing the glutathione S-transferase
(GST)-C21 construct (a kind gift of Dr. J. G. Collard) that
encodes the Rho-binding domain of Rhotekin (a Rho effector protein)
fused to GST. GST-C21 protein synthesis was induced with 0.1 mM
isopropyl -D-thiogalactoside. After 12 h, the
culture was centrifuged (3 900 g, 20 min, 4°C), and the
pellet was sonicated in a bacterial lysis buffer [50 mM
Tris · HCl, pH 7.4, 100 mM NaCl, 1 mM EDTA, 5% glycerol, 0.1%
Triton X-100, 2 mM dithiothreitol (DTT), 1 mM phenylmethylsulfonyl
fluoride (PMSF), 0.4 mM sodium pervanadate, 10 µg/ml leupeptin, and
10 µg/ml aprotinin]. The lysate was centrifuged (13 000 g, 20 min, 4°C), and the supernatant containing the
GST-C21 fusion protein was incubated with 50% gluthatione-Sepharose 4B beads (Amersham) during 60 min at 4°C. The beads were washed three times with and resuspended in GST-fish buffer (50 mM
Tris · HCl, pH 7.4, 100 mM NaCl, 2 mM MgCl2, 10%
glycerol, 1% Nonidet P-40, 2 mM DTT, 1 mM PMSF, 0.4 mM sodium
pervanadate, 10 µg/ml leupeptin, and 10 µg/ml aprotinin).
Visualization of F-actin fibers.
CPAE cells (either nontransfected cells stimulated with 25% HTS or 2.5 U/ml thrombin or transfected cells expressing the constitutively active
forms of G13, RhoA, or Rho kinase) were fixed for 10 min at room temperature in 3.7% formaldehyde in PBS and permeabilized in
0.2% Triton X-100 in PBS for 4 min. After being blocked with 10% FCS
in PBS for 30 min, cells were incubated with 1 unit of rhodamine-phalloidin (Molecular Probes) diluted in PBS with 1% FCS per
coverslip for 20 min at room temperature. Cells were rinsed twice in
PBS after each incubation. Finally, the cells were washed twice with
water. To prevent photobleaching, cells were mounted in Vectashield
(Vector Laboratories). Labeled cells were observed with 1) a
Nikon Optiphot-2 microscope using green excitation for visualization of
rhodamine staining and blue excitation to identify the green
fluorescent cells, or 2) a Leica SP-2 laser scan confocal microscope.
Quantification of F-actin. The cellular F-actin content was estimated essentially as described previously (27), except that cells were attached to multiwell plastic dishes. Cells were seeded at 90,000 cells/well and after stimulation with 25% HTS or 2.5 U/ml thrombin fixed in 3.7% paraformaldehyde in TBS. The cells were washed in 0.1% saponin buffer (in mM: 10 MOPS pH 6.9, 5.5 EGTA, 20 K2HPO4, 1.95 MgSO4, and 0.1% wt/vol saponin) for permeabilization and incubated with 200 µl of 0.33 µM rhodamine-phalloidin diluted in 0.1% saponin buffer. After 1 h at room temperature, cells were washed twice in MOPS buffer (in mM: 10 MOPS pH 6.9, 5.5 EGTA, 20 K2HPO4, and 1.95 MgSO4) and incubated in 1 ml methanol for 30 min to extract the rhodamine-phalloidin. The solution was transferred to a cuvette, and methanol was added to a final volume of 2.5 ml. Rhodamine fluorescence was measured spectrofluorometrically at 576 nm after excitation at 540 nm using a PTI RatioMaster spectrophotometer. As a control, identical samples were incubated as described above, except in the presence of a 100-fold excess of unlabeled phalloidin (Molecular Probes). The fluorescence measured from these samples was <4% of that obtained under standard conditions. Data for swollen and thrombin-stimulated cells are presented as the 576-nm emission (after subtraction of a methanol blank) normalized to the blank-subtracted rhodamine signal of nonstimulated control cells (relative scale). The assay was verified to be linear over a range of at least 60,000-100,000 cells per well (data not shown).
Biochemical effect of C2IN-C3 toxin on Caco-2 cells. Caco-2 cells were incubated with C2IN-C3 toxin by adding 200 ng C2II and 200 ng C2IN-C3/ml culture medium. After overnight incubation, cells were lysed in a 1% Triton buffer containing (in mM) 25 Tris, 100 NaCl, 90 mannitol, 2 DTT, 1 EGTA, and 1 PMSF, as well as 10 µg/ml aprotinin and 10 µg/ml leupeptin. The lysate was centrifuged (500 g, 4°C, 5 min), and aliquots of the supernatant were used for 12% SDS-PAGE. After blotting, Rho was detected with a monoclonal antibody (Santa Cruz Biotechnology sc-4178; diluted 1:2,000) and a secondary alkaline-phosphatase-conjugated goat anti-mouse IgG (diluted 1:7,000) as described previously (26).
Data analysis. Analysis of electrophysiological data was performed using the WinASCD software (Guy Droogmans, Laboratorium voor Fysiologie, KU Leuven). The Origin software package version 6.0 (Microcal Software, Northampton, MA) was used for statistical analysis and graphical presentation of the data.
Time courses of the whole cell current were obtained by plotting the current at ![]() |
RESULTS |
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Transient expression of constitutively active G13,
RhoA, or Rho kinase does not affect ICl,swell in CPAE
cells.
In a first series of experiments, we investigated the effect of
constitutively active isoforms of G
13 (mutation
Gln226Leu; see Ref. 16), RhoA (mutation Gln63Glu; see Ref.
30) or Rho kinase (CAT: Rho kinase containing only the
catalytic domain; see Ref. 2) on
ICl,swell in transiently transfected CPAE cells. Under isotonic conditions, the basal membrane current of control CPAE
cells had a mean density of 13.4 ± 2.8 pA/pF at +100 mV
(n = 31; Fig. 1). CPAE
cells transfected with RhoA Gln63Glu developed slightly larger basal
membrane currents during isotonicity with a mean density of 26 ± 5.3 pA/pF (n = 15) at +100 mV (Fig. 1). However, this
value was not significantly different from that in control cells.
Similarly, the basal membrane current in CPAE cells transfected with
either G
13 Gln226Leu or Rho kinase-CAT did not differ
significantly from that in wild-type CPAE cells: the basal membrane
current had a mean density of, respectively, 18.4 ± 3.4 pA/pF
(n = 40) and 16.8 ± 5.54 pA/pF (n = 16) at +100 mV (Fig. 1).
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Hypotonic cell swelling by 25% HTS does not activate the Rho
pathway in CPAE cells.
In a complementary series of experiments, we investigated whether
hypotonic cell swelling triggers activation of the Rho pathway. To
measure Rho activation, we used a pull-down assay in which the active
Rho-GTP, but not the inactive Rho-GDP, is bound to a GST fusion protein
containing the Rho binding domain of Rhotekin (39).
Rho-GTP formation was monitored during a 4-min 25% HTS, which in CPAE
cells is sufficiently long to fully activate VRACs, since we have
previously shown that it takes between 60 and 90 s for a 25% HTS
to half-maximally activate VRACs in CPAE cells (25, 36).
As shown in Fig. 3, the ratio of active
Rho-GTP to total Rho did not change significantly during hypotonic
swelling of CPAE cells. In contrast, stimulation of CPAE cells with
thrombin (5 U/ml) induced the formation of Rho-GTP (Fig. 3), as has
also been observed for human umbilical vein endothelial cells
(39).
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The Rho pathway is not required for activation of VRACs in
caveolin-1 transfected Caco-2 cells.
In a second part, we addressed the question whether a functional Rho
pathway is a general requirement for VRAC activation as is observed in
CPAE cells (25, 26) or, alternatively, whether VRAC
activation can occur independently of the Rho pathway. These experiments were performed in Caco-2, a colon carcinoma cell line that
does not express endogenous caveolin-1 (37). We have
previously shown that a hypotonic stimulus is unable to activate VRACs
in wild-type Caco-2 cells, but that expression of caveolin-1 restores efficient VRAC activation (37). We first confirmed by
Western blotting that RhoA is expressed in Caco-2 cells (data not
shown; see also Fig. 6D, inset), as has also been
demonstrated by others (11). Subsequently, we investigated
whether manipulating the Rho pathway, either by stimulation with
GTPS or by inhibition with the C2IN-C3 fusion toxin, modulated
ICl,swell in wild-type and caveolin-1
transfected Caco-2 cells.
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DISCUSSION |
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We have previously shown that activation of VRACs in vascular
endothelial cells (CPAE) requires a functional Rho/Rho kinase/myosin light chain phosphorylation pathway, since inhibition of Rho, Rho
kinase, or myosin light chain kinase abrogates the swelling-induced activation of VRACs. Yet, in this study we show that activation of the
Rho pathway by constitutively active isoforms of G13, Rho, or Rho kinase has no effect on VRACs either in isotonic or in
hypotonic conditions. Importantly, we did not observe spontaneous activation of VRACs in control conditions or potentiation during cell
swelling. In a complementary series of experiments we examined whether
cell swelling would activate the Rho pathway, but there was no
indication that this pathway was turned on by cell swelling. These data
are consistent with previous observations on the effect of thrombin on
VRACs: thrombin is unable to trigger VRACs under isotonic conditions
(21), despite it being a potent stimulator of the Rho
pathway in endothelial cells (9, 39). In view of the
inability of the Rho pathway to activate VRACs and the lack of
activation of this pathway during hypotonic cell swelling, we conclude
that the Rho pathway does not transmit the activator signal that opens
VRACs during hypotonic cell swelling. At first sight, this conclusion
may seem to contradict previous observations in CPAE cells that GTP
S
activates VRACs under isotonic conditions and that inhibition of the
Rho pathway precludes this GTP
S effect (25, 41).
However, one should bear in mind that the GTP
S effect is transient
with VRAC activity being restored to baseline level within 5-10
min of GTP
S dialysis. Moreover, after the initial transient phase,
GTP
S-treated cells and control cells behave identically when exposed
to a hypotonic stimulus. Thus, in the longer term, GTP
S does not
affect VRAC activation, which is consistent with the lack of effect of
constitutively active G
13, RhoA, or Rho kinase on VRAC activity.
The conclusion that the Rho pathway does not play a causal role in swelling-induced activation of VRACs can be reconciled with the previously established requirement for a functional Rho pathway by postulating a permissive effect for the Rho pathway, i.e., Rho pathway activity is required for but it does not trigger opening of VRACs during hypotonic cell swelling. One implication of such a permissive effect would be that the Rho pathway could help to sensitize the cell to hypotonic stimuli. Indeed, we have previously observed that thrombin potentiates VRACs at mild hypotonic stimulation (13%) but less so at stronger hypotonic stimuli (28% HTS) (21). In addition, it would explain why thrombin or a myosin light chain phosphatase inhibitory peptide (NIPP1191-210) potentiates VRACs once the channels have been preactivated by a hypotonic stimulus, although they exert no effect on VRACs under isotonic conditions (21, 25). What could be the molecular basis for such a permissive effect? As reviewed by Janmey (14), the cytoskeleton forms a three-dimensional network that not only determines the mechanical properties of the cell but also forms an extended scaffold onto which regulatory and signaling proteins can bind. In this context, the Rho pathway could facilitate VRAC activation by promoting the formation of actin filaments that would serve as a platform on which proteins that participate in the VRAC activation cascade assemble.
Although we were unable to detect alterations in Rho activity, F-actin stress fibers, or net cellular F-actin content during cell swelling, cytoskeletal rearrangements during hypotonic swelling have been described before (17, 19, 23, 27, 35). Pedersen et al. (27) and Levitan et al. (19) reported a disruption of the cortical F-actin cytoskeleton in, respectively, Ehrlich ascites tumor cells and B-lymphocytes, whereas formation of membrane ruffles due to cortical actin polymerization was observed in intestinal 407 cells (35) and C6 glioma cells (23). Koyama et al. (17) observed a transient formation of F-actin stress fibers during hypotonic swelling of bovine aortic endothelial cells. However, it was concluded that the observed cytoskeletal rearrangements during hypotonic cell swelling were not critical in initiating regulatory transport processes (17, 19, 23).
A second finding of this study is the variable requirement for a
functional Rho pathway. In CPAE cells, intracellular application of
GTPS triggers a transient outwardly rectifying chloride current that
is phenotypically identical to ICl,swell
(41). This effect of GTP
S is mediated via Rho and Rho
kinase, since inhibition of Rho with the C3 exoenzyme or of Rho kinase
with Y-27632 abolishes the GTP
S-induced current in endothelial cells
(26). More recently, Koyama et al. (17)
showed that inhibition of Rho with C3 toxin or of Rho kinase with
Y-27632 also reduced the swelling-induced ATP release by bovine aortic
endothelial cells. Thus, in vascular endothelial cells,
swelling-induced responses seem critically dependent on a functional
Rho system. A similar conclusion can be drawn for N1E115 neuroblastoma
cells in which blockade of Rho by the C. difficile toxin B
also reduces ICl,swell (10). In contrast, caveolin-1-expressing Caco-2 cells were unresponsive to
GTP
S despite normal VRAC activation by a hypotonic stimulus. Furthermore, C3 exoenzyme pretreatment did not affect the HTS-triggered activation of VRACs. First, this is consistent with the previous conclusion that the Rho pathway does not transmit the primary activating signal during cell swelling. Second, it also suggests that
the permissive effect of the Rho pathway is either dispensable in
Caco-2 cells and/or that there are alternative structures that can take
over the permissive role of the Rho pathway. This conclusion is
compatible with the previously reported failure of GTP
S to activate
ICl,swell in Xenopus oocytes
(1) or in C6 glioma cells (34). It therefore
seems that, at least in these experimental systems, Rho is not required
for swelling-induced activation of VRACs.
In summary, the present data, in combination with our previous observations on VRACs in CPAE cells, allow us to delineate more precisely the contribution of the Rho pathway to hypotonicity-induced activation of VRACs. An intact Rho/Rho kinase/myosin light chain phosphorylation pathway is required for VRAC activation in CPAE cells. However, it does not transmit the activator signal that opens VRACs during cell swelling. We therefore propose that the Rho pathway fulfils a permissive role, but not a causal role, during VRAC activation in CPAE cells.
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ACKNOWLEDGEMENTS |
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The technical help of J. Prenen, A. Florizoone, M. Crabbé,
and H. Van Weijenbergh is greatly appreciated. We thank J. Collard (The
Netherlands Cancer Institute, Amsterdam, The Netherlands) and M. Schwartz (The Scripps Research Institute, La Jolla, CA) for providing
the GST-C21 and the GST-RBD expression vector; M. Negishi (Laboratory
of Molecular Neurobiology, Kyoto University, Japan) for the
G13Gln226Leu cDNA; and K. Kaibuchi (Nara Institute of
Science and Technology, Nara, Japan) for the Rho kinase-CAT construct.
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FOOTNOTES |
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This research was supported by grants of Interuniversitaire Attractiepolen (IUAP P4/23), FWO-Vlaanderen (FWO G.0214.99), Geconcerteerde Onderzoeksactie (GOA 99/07), and the Danish Natural Science Research Council.
Address for reprint requests and other correspondence: J. Eggermont, Laboratorium voor Fysiologie, KU Leuven, Campus Gasthuisberg, B-3000 Leuven, Belgium (E-mail: Jan.Eggermont{at}med.kuleuven.ac.be).
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 February 20, 2002;10.1152/ajpcell.00038.2001
Received 25 January 2001; accepted in final form 18 February 2002.
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