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The Journal of Neuroscience, February 15, 1998, 18(4):1196-1206
Mitogen-Activated Protein and Tyrosine Kinases in the Activation
of Astrocyte Volume-Activated Chloride Current
Valérie
Crépel1,
William
Panenka1,
Melanie E. M.
Kelly2, and
Brian A.
MacVicar1
1 Neuroscience Research Group, University of Calgary,
Calgary, Alberta, Canada T2N 4N1, and 2 Department of
Pharmacology, Dalhousie University, Halifax, Nova Scotia B3H 4H7
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ABSTRACT |
Astrocytes swell during neuronal activity as they accumulate
K+ to buffer the increase in external
K+ released from neurons. This swelling activates
volume-sensitive Cl channels, which are thought to
be important in regulatory volume decrease and in the response of the
CNS to trauma and excitotoxicity. Mitogen-activated protein (MAP)
kinases also are activated by cell volume changes, but their roles in
volume regulation are unknown. We have investigated the role of
tyrosine and MAP kinases in the activation of volume-activated
Cl channels in cultured astrocytes, using
whole-cell patch-clamp recording and Western immunoblots. As previously
described, hypo-osmotic solution induced an outwardly rectifying
Cl current, which was blocked by NPPB and SITS.
This Cl current did not depend on
[Ca2+ ]i because it was still observed
when 20 mM BAPTA was included in the pipette, but it did
exhibit rundown when ATP was omitted. Inhibition of tyrosine kinases
with genistein or tyrphostin A23 (but not the inactive agents daidzein
and tyrphostin A1) blocked the Cl current. The MAP
kinase kinase (MEK) inhibitor PD 98059 reversibly inhibited activation
of the Cl current by hypo-osmotic solution.
Western immunoblots showed that genistein or PD 98059 blocked
activation of Erk-1 and Erk-2 by hypo-osmotic solution in astrocytes.
Therefore, activation of tyrosine and MAP kinases by swelling is a
critical step in the opening of volume-sensitive
Cl channels.
Key words:
volume-activated Cl current; tyrosine kinase; MAP kinase; cell swelling; regulatory volume decrease; astrocyte
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INTRODUCTION |
The swelling of astrocytes is an
important response to neuronal activity in the CNS and occurs during
normal and pathological conditions (for review, see Kimelberg, 1995 ).
Astrocytes are thought to be the most labile in generating cellular
volume changes. During neuronal activity there is an increase in
extracellular K+, which is taken up into astrocytes
via transport and passive ion flows. This, in turn, causes a net uptake
of water and swelling in astrocytes and a reduction of the
extracellular space (for review, see Kimelberg, 1995). Astrocytes and
other cell types counteract volume increases by an efflux of
Cl and other anions, coupled with the coactivation
of K+ channels (Pasantes-Morales et al., 1994) (for
review, see Nilius et al., 1996; Strange et al., 1996). The
Cl channel involved in volume regulation is
outwardly rectifying and is permeable to fairly large anions, including
glutamate and taurine. However, the cellular mechanisms controlling the
activation of the volume-sensitive Cl channels in
astrocytes are poorly understood.
The mitogen-activated protein (MAP) kinases Erk-1 and Erk-2 can be
activated by hypo-osmotic solution and swelling (Tilly et al., 1993;
Schliess et al., 1995, 1996; Noé et al., 1996) (but see Krause et
al., 1996). Growth factors, cytokines, and some G-protein-coupled
receptors activate MAP kinases, which are involved in the regulation of
gene expression (for review, see Su and Karin, 1996; Robinson and Cobb,
1997). There are also several cytoplasmic targets that are modulated by
MAP kinases (Campbell et al., 1995). However, the roles for MAP kinases
in regulating ion channel activity are unknown. In contrast, it has
been shown that tyrosine kinases are involved in the activation of the
volume-sensitive Cl channels in Intestine 407 and
in cardiac cells (Tilly et al., 1993; Sorota, 1995). Because MAP
kinases can be targets of tyrosine kinase cascades, we suggest that
both tyrosine and MAP kinases may be involved in the activation of
volume-sensitive Cl channels in astrocytes.
In this paper we investigated the involvement of tyrosine kinase and
the MAP kinases, Erk-1 and Erk-2, in the activation of a
volume-sensitive Cl current in cultured
astrocytes, using whole-cell patch-clamp recording and Western
immunoblots. Specific inhibitors of tyrosine kinases (genistein and
tyrphostin A23; Negrescu et al., 1995) (for review, see Levitzki and
Gazit, 1995) were used, and their specificity was tested by using the
inactive structural analogs of these compounds (daidzein for genistein
and tyrphostin A1 for tyrphostin A23). To prevent Erk-1 and Erk-2
activation, we used the specific MAP kinase kinase (MEK) inhibitor PD
98059 (Dudley et al., 1995). We report that hypo-osmotic solution
induced an outwardly rectifying Cl current, which
was [Ca2+]i-independent and
[ATP]i-dependent. Tyrosine kinase and MEK inhibitors reversibly blocked activation of volume-sensitive
Cl currents. In addition, Western immunoblots
showed that activation of Erk-1 and Erk-2 by hypo-osmotic solution
depended on the activation of tyrosine kinases and MEK. In conclusion,
the volume-sensitive Cl channel appears to be a
cytoplasmic target for the MAP kinase signaling pathway.
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MATERIALS AND METHODS |
Cell culture. Astrocyte cultures were prepared from
1-d-old Sprague Dawley rats (University of Calgary, Alberta, Canada), using modifications of standard techniques (McCarthy and Devellis, 1980; Merrill et al., 1984). All procedures conformed to guidelines laid down by the Canadian Council on Animal Care. Briefly, neonates were decapitated and the brain exposed. The cortex was dissected free
of the underlying brain tissue under sterile conditions, and the
meninges and pia mater were removed. The tissue was dissociated by
mechanical trituration, and the resulting cell suspension was plated
onto glass coverslips and grown in DMEM and Ham's F12 (1:1) with 10%
fetal calf serum. Medium was changed twice per week. As previously
described (MacVicar et al., 1991), immunocytochemical characterization
of cultures indicated that >95% of cells stained positive for glial
fibrillary acidic protein, confirming that the primary cell type
present is astrocytes (data not shown).
Recording procedures. Coverslips with adherent glial cells
were placed in a recording chamber (volume, ~200 µl) and mounted on
an inverted microscope (Zeiss, Oberkochen, Germany). Cells were
observed via phase-contrast optics and superfused at 1-2 ml/min
(20-22°C). Standard extracellular salt solution for recording anion
currents was composed of (in mM): 70 Trizma-HCl, 100 sucrose, 1.5 CaCl2, 10 HEPES, and 10 glucose,
adjusted to pH 7.3 with CsOH. TEA (5 mM) and
BaCl2 (5 mM) were added to the extracellular
solution to eliminate K+ currents. Solution
osmolarity was determined by a vapor pressure osmometer (5120C, Wescor,
Logan, UT). The extracellular solution was between 280 and 290 mOsm/l.
In experiments testing hypo-osmotic stimulation (HOS), the sucrose was
removed from the bathing solution (220 mOsm/l). We observed that, in
this hypo-osmotic condition, astrocytes swelled as previously reported
(Pasantes-Morales et al., 1994; Lascola and Kraig, 1996).
Ionic currents were recorded with the patch-clamp technique. Patch
pipettes were pulled from borosilicate thin-wall glass capillaries
(TW150F-4, World Precision Instruments, Sarasota, FL) with the
Flaming/Brown micropipette puller (P-97, Sutter Instrument, Novato, CA)
and had tip resistances of 3-7 M when filled with electrode
solution. The standard pipette solution was composed of (in
mM): 60 Trizma-HCl, 70 Trizma-base, 70 aspartic acid, 15 HEPES, 0.4 CaCl2, 1 MgCl2, 1 ATP,
0.5 GTP, and 1 EGTA, adjusted to pH 7.25 with CsOH. In some other
experiments the intracellular concentration of Cl
was decreased by replacing 30 mM Trizma-HCl by a 30 mM concentration of the less-permeant anion aspartate
(Lewis et al., 1993) in the electrode solution. Free
Ca2+ in the standard pipette solution was estimated
to be ~100 nM. In those experiments in which
Ca2+ was chelated by using 20 mM BAPTA
in the pipette to replace EGTA, free Ca2+ was
calculated to be reduced to <10 nM. To investigate current decay, we performed some experiments in the absence of ATP and/or GTP
in the electrode solution.
Membrane currents were recorded in whole-cell configuration in the
voltage-clamp mode (VH =  60 mV) with an
Axopatch-1D amplifier (Axon Instruments, Foster City, CA). Currents
were filtered with a four-pole low-pass Bessel filter (3 dB at 5 kHz). Offset potentials were nulled, using the amplifier circuitry
before seals were made on cells. Liquid junction potentials (LJP)
between the bath and patch-clamp electrodes were measured
experimentally and defined as the potential of the bath with respect to
the electrode solution (Barry and Lynch, 1991). For whole-cell
recording the membrane potential of the cell,
Vm, was calculated as:
Vm = Ve LJP. When the extracellular solution contained 88 mM Cl and the intracellular solution
contained 63 or 33 mM Cl, the LJP was
1 or 2-3 mV, respectively. The membrane potential was not corrected
for the LJP, because the variation of potential was minor.
Cell capacitance and series resistance values were measured and
compensated by using the circuitry of the amplifier. The intrinsic membrane input resistance of the cell was monitored by the application of hyperpolarizing voltage steps of 10 mV (500 msec duration) within
the initial 5-10 min of the experiment. The voltage dependence of
ionic currents was studied by using voltage ramp commands: the membrane
potential was stepped from VH = 60 to 120
mV, held at 120 mV for 50 msec, and then ramped to + 80 mV in 2 sec.
The efficacy of the voltage clamp during the ramp command was tested by
comparing the I/V curve obtained with the voltage ramp with that obtained with voltage steps (10 mV amplitude increments and 500 msec duration). Because no difference was found between I/V curves obtained by using either ramp or step commands over the voltage
range tested (120 to +80 mV), only I/V curves obtained with the voltage ramp commands are shown in this report.
Data analysis. Membrane currents were digitized every 20 sec
with pClamp software (Axon Instruments). Continuous recordings of
current and voltage were displayed on a chart recorder (RS 3200, Gould,
Cleveland, OH). Data were analyzed off-line and are presented as
mean ± SEM. Statistical analyses were performed with Student's
paired t test, and data were considered significantly different at p < 0.05.
Western immunoblotting. Confluent astrocyte cultures were
starved in serum-free DMEM for 24-30 hr. After this period the
cultures were transferred to a defined medium containing (in
mM) 110 NaCl, 58 NaCO3, 25 glucose, 20 HEPES, 5 KCl, 0.6 MgSO4, 0.9 NaHPO4, and 1.8 CaCl2 for a further 8 hr. Hypo-osmotic treatment consisted of lowering the extracellular NaCl
concentration from 110 to 33 mM by adding NaCl-free medium
to the cultures. This represented a drop in osmolarity of ~30%, as
measured by vapor pressure osmometry. For the iso-osmotic condition,
NaCl-free medium was supplemented with mannitol to maintain original
osmolarity while still controlling for the drop in salt concentration.
After stimulation with hypo-osmotic solution, cultures were washed with
ice-cold PBS and harvested on ice with protein sample buffer (10%
glycerol, 62.5 mM Tris-base, 2% SDS, and 2.5%
 -mercaptoethanol). Then the samples were boiled, sonicated briefly,
and subjected to SDS-Page (Laemmli, 1970) on 15% w/v acrylamide gels.
The protein was transferred to polyvinylidene fluoride (PVDF) at 70 mA
overnight. After transfer, the membranes were blocked in TTBS (0.05%
Tween 20, 25 mM Tris-base, and 62.5 mM NaCl,
solution pH-adjusted to 7.5) containing 5% w/v skim milk and then
incubated in primary antibody. For MAP kinase blots, polyclonal
anti-rat MAP kinase R2 (Erk-1-CT; 1:2000 dilution) was used in TTBS
with 3% nonfat dry milk; the incubation time was ~2 hr at room
temperature. The monoclonal anti-phosphotyrosine antibody 4G10 was used
for phosphotyrosine blots at a dilution of 1:1000 in 1% BSA in TTBS
and incubated overnight at room temperature.
After being incubated with the primary antibodies, membranes were
washed and incubated in secondary antibodies. For blots in which
polyclonal primary antibodies were used, the secondary antibody was an
anti-rabbit IgG, horseradish peroxidase-linked (H+L) whole antibody
(1:3000). For monoclonal primary antibodies, the secondary antibody was
goat anti-mouse IgG (H+L) diluted at 1:5000. All secondary antibodies
were diluted in the same buffer as their respective primary antibodies,
and membranes were incubated in the secondary antibodies for 2 hr at
room temperature.
After secondary antibody incubation, membranes were washed and probed
with the Supersignal Substrate Western Blotting ECL detection
kit.
Materials. All culture reagents were purchased from Life
Technologies (Burlington, Ontario). Genistein, daidzein, tyrphostin A23, tyrphostin A1, and PD 98059 were purchased from Calbiochem (La
Jolla, CA). 5-Nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) was
purchased from BIOMOL Research Laboratories (Plymouth Meeting, PA).
4-Acetamido-4"-isothiocyanatostilbene-2,2'-disulfonic acid (SITS) and
all other drugs were purchased from SIGMA (Oakville, Ontario). Primary
antibodies for Western blotting were obtained from Upstate
Biotechnology (Lake Placid, NY). Secondary antibodies were obtained
from Amersham (Oakville, Ontario; anti-rabbit IgG) and Bio-Rad
laboratories (Mississauga, Ontario; goat anti-mouse IgG).
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RESULTS |
The recordings were performed on flat polygonal or stellate
astrocytes. After the whole-cell recording configuration was
established, the membrane current was recorded in voltage-clamp mode at
a holding potential of 60 mV. Data presented here were selected from
cells that had a mean membrane capacitance and input resistance of
40.4 ± 3.7 pF (n = 35) and 2.32 ± 0.4 G
(n = 35), respectively.
Hypo-osmotic stimulation activates a Cl
current in cultured astrocytes
As shown in Figure
1A, a short (3-5 min)
superfusion of cells with HOS resulted in the reversible activation of
an inward current at 60 mV. The voltage dependence of HOS-induced
current was studied by plotting I/V relations that were
derived with a 2 sec depolarizing ramp command from 120 to +80 mV.
Figure 1B shows the I/V plots for currents
recorded before (1), during (2), and after
(3) recovery from HOS exposure. The I/V curve of
HOS-induced current was obtained by subtracting the peak current
recorded during HOS application from that recorded before HOS
application (Fig. 1C). The HOS-induced current was outwardly
rectifying and reversed at approximately 10 mV. The slope conductance
calculated for two representative membrane potential intervals, 120
to 60 mV (referred to as negative conductance) and +20 to +80 mV
(referred to as positive conductance), was 5.26 ± 0.52 nS
(n = 35) and 15.3 ± 1.29 nS (n = 35), respectively.
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Figure 1.
Hypo-osmotic stimulation activates an outwardly
rectifying Cl current in cultured astrocytes.
A, Membrane current recorded in response to voltage ramp
commands (from 120 to + 80 mV, 2 sec duration, 0.05 Hz) before and
during two successive hypo-osmotic (HOS) stimulations (5 min duration, VH = 60 mV) in a cultured astrocyte with (in mM) 63 Cl, 2 ATP,
and 0.5 GTP in the pipette solution. In this and the following figures
the top and the bottom traces illustrate
the current and the voltage traces, respectively. B,
Individual current-voltage (I/V) relations
obtained at the times indicated by the numbers in
A. C, Mean I/V relations
of HOS-induced current obtained by subtracting the peak current
recorded during HOS application (2) from that
recorded before HOS application (1)
(n = 35). D, Mean I/V
relations of HOS-induced current obtained with either 33 mM Cl ( , n = 9) or 63 mM Cl ( , n = 14) in the pipette solution (minus ATP and GTP). Note that HOS-induced
current was outwardly rectifying and reversed at approximately 25 mV
(downward arrow, D) or 10 mV
(upward arrow, D) with either 33 or 63 mM Cl in the pipette solution,
respectively. In this and the following figures, the error bars
represent the SEM.
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To characterize the ionic selectivity of HOS-induced current, we
examined the change of the reversal potential of this current with
alterations in the transmembrane Cl gradient.
Cells were held at 60 mV and ramped from 120 to +80 mV in standard
extracellular Ringer's ([Cl]out = 88 mM)
with either 33 or 63 mM [Cl] in the
electrode solution (and no ATP and GTP). A reduction in
[Cl]in shifted the reversal
potential of HOS-induced current to more negative potentials. Figure
1D shows that the reversal potential of the current
was 9.5 ± 0.8 mV (n = 14) in 63 mM
Cl and 24 ± 1.6 mV (n = 9)
in 33 mM [Cl]i. These
values are nearly identical to those predicted by the Nernst equation
(8.6 and 25.4 with [Cl]in = 63 and 33 mM, respectively) and confirm that the HOS-induced current is carried mainly by Cl in our
experimental conditions.
To characterize HOS-induced current further, we tested the effect of
the Cl channel blockers NPPB and SITS on this
current. As shown in Figure 2A, a 3 min external
application of either NPPB (0.1 mM) or SITS (0.5 mM) rapidly and reversibly decreased HOS-induced current. Figure 2, B and C, shows the I/V
relations recorded for HOS-induced current in the absence
(Control) and presence of either NPBB or SITS. NPPB
reduced HOS-induced current over the full range of potentials tested,
with a similar decrease in both the negative conductance (by 89 ± 7.5%; p = 0.026; n = 5) and the
positive conductance (by 93 ± 5.2%; p = 0.018;
n = 5). The SITS block of the HOS-induced current, in
contrast, was strongly voltage-dependent, as previously described
(Kelly et al., 1994; Arreola et al., 1995). In the presence of SITS the
positive HOS conductance was reduced by 94.3 ± 2.75%
(p < 0.001; n = 11), but the
negative conductance was not significantly modified (0.5 ± 5.9%; p = 0.75; n = 11). Thus, under
our experimental conditions, hypo-osmotic solution, which induces cell
swelling, activates a Cl current that will be
referred to hereafter as volume-activated Cl
current or ICl,vol, as previously named
in the literature (for review, see Nilius et al., 1996).
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Figure 2.
ICl,vol is blocked by
NPPB and SITS. A, Membrane current and conductance
changes evoked by a long HOS application (23 min in duration,
VH = 60 mV) recorded in the absence or in
the presence of either NPPB (0.1 mM) or SITS (0.5 mM) in the external media [and (in mM) 63 Cl, 2 ATP, and 0.5 GTP in the pipette solution].
B, C, Mean I/V relations
of HOS-induced current obtained before (), during ( ), and after
(+) bath application of either NPPB (B;
n = 5) or SITS (C;
n = 11). Note that HOS-induced current was strongly
reduced by NPPB in a voltage-independent manner and by SITS in a
voltage-dependent manner.
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Activation of ICl,vol does not require a
change in intracellular Ca2+
Because cell swelling induced by a hypo-osmotic shock is known to
be associated with an increase of internal Ca2+
([Ca2+]i) in some cell types
(for review, see Hoffmann, 1992), we studied the dependence of
ICl,vol on
[Ca2+]i. For these experiments
ICl,vol was recorded in the presence of the
Ca2+ chelator BAPTA (20 mM) in the
pipette solution and a low external concentration of
Ca2+ (0.15 mM) to prevent a rise of
[Ca2+]in during the cell swelling.
Figure 3A shows whole-cell
currents recorded from a representative cell exposed to a 3 min
application of HOS in low [Ca2+]out
with 20 mM BAPTA/63 mM
[Cl] in the electrode. HOS still resulted in
reversible activation of ICl,vol under these
conditions. Figure 3B shows the mean I/V curve
obtained for the peak current activated by HOS in five cells. Current
amplitudes were comparable when recorded in either a low external
concentration of Ca2+ with BAPTA in the pipette
solution (67 ± 14.5 pA/pF at +80 mV, n = 5) or in
standard Ca2+ containing external solutions with no
BAPTA in the electrode solution (44 ± 6.4 pA/pF at +80 mV,
n = 27). These data confirm that the activation of
ICl,vol is independent of an increase in [Ca2+]i, as previously
described in other systems (Hagiwara et al., 1992; Nilius et al., 1996)
(for review, see Strange et al., 1996).
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Figure 3.
Activation of ICl,vol
is independent of a change in intracellular Ca2+.
A, Membrane current evoked by an HOS application (3 min
duration, VH = 60 mV) recorded in the
presence of the Ca2+ chelator BAPTA (20 mM) with (in mM) 63 Cl, 2 ATP, and 0.5 GTP in the pipette solution. The external concentration of
Ca2+ was reduced to 0.15 mM.
B, Mean I/V relations of
ICl,vol recorded in five separate cells
under the same conditions as for currents shown in A
(n = 5). Note that, in the presence of the
Ca2+ chelator BAPTA in the pipette solution and a
low external concentration of Ca2+, the HOS
stimulation still evoked an outwardly rectifying current that reversed
at approximately 10 mV.
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Activation of ICl,vol depends on the
cellular metabolic state
We next investigated other possible mechanisms involved in the
regulation of ICl,vol. Studies performed in
kidney, T lymphocytes, and endothelial cells have shown that activation
of volume-sensitive anion currents depends on the metabolic state of
the cell and is linked to cellular nucleotide levels (Lewis et al.,
1993; Jackson et al., 1994; Oike et al., 1994; Strange et al., 1996).
To test this hypothesis in astrocytes, we compared
ICl,vol in the presence and the absence of ATP
and GTP in the pipette solution (Fig. 4). We showed that under control conditions, with 2 mM ATP and
0.5 mM GTP in the pipette solution (Fig.
4A, 1), a long exposure to HOS (at least
30 min) resulted in the activation of a stable
ICl,vol, which reached a maximal
amplitude of 45.4 ± 9.2 pA/pF (at +80 mV, n = 6)
in 16 ± 3.5 min (n = 6) and showed no decline
over the period of HOS exposure. In contrast,
ICl,vol, recorded during 30 min of HOS in
the absence of ATP and GTP in the pipette solution (Fig.
4A, 2), reached a maximal amplitude of
16.3 ± 2.9 pA/pF (at +80 mV, n = 10) in 7.8 ± 0.72 min (n = 10) but then progressively declined at
a rate of 2.25 ± 0.2% per minute (n = 10). In
the absence of ATP and GTP the current amplitude ran down by one-half in 16.5 ± 2.4 min (n = 10), and the maximal
amplitude of ICl,vol (16.33 ± 2.9 pA/pF at
+80 mV, n = 10) was significantly smaller than that
recorded under control conditions (45.4 ± 9.2 pA/pF at +80 mV;
n = 6; p < 0.0025). To discriminate
between ATP binding and hydrolysis, we recorded the rundown of
ICl,vol while including AMP-PNP, a
nonhydrolyzable form of ATP, in the pipette solution. GTP also was
included in the pipette. In these conditions the rundown of
ICl,vol still occurred, indicating that
hydrolysis of ATP was necessary (n = 10; data not
shown). Therefore, the activation of ICl,vol
apparently involves phosphorylation processes.
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Figure 4.
ICl,vol is regulated by
the cellular nucleotides. A, Membrane current evoked by
a long HOS application (30 min in duration, VH = 60 mV) recorded in the presence
(1) or in the absence (2) of ATP (2 mM) and GTP (0.5 mM) in the pipette
solution. B-D, Graphs showing the changes of the
normalized holding current (B), negative conductance (C), and positive conductance
(D) of ICl,vol versus time during the HOS application (time 0 corresponds to
the beginning of the HOS application), either in the presence
(1; n = 6) or in the absence
(2; n = 10) of ATP and GTP in the
pipette solution. The positive and negative conductances were
calculated for potentials ranging from +20 to +80 mV and 120 to 60
mV, respectively. The holding current and the negative and positive
conductances have been normalized to 1, using their maximal values
obtained during the HOS application. Note that activation of
ICl,vol was stable in the presence of ATP
and GTP. In contrast, when ATP and GTP were omitted from the pipette
solution, ICl,vol progressively declined.
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Tyrosine and MAP kinases are involved in the regulation of
ICl,vol
Previous studies on other cell types have shown that
volume-activated Cl currents can be modulated by
tyrosine kinase-dependent phosphorylation (Tilly et al., 1993; Sorota,
1995; Strange and Jackson, 1995). To investigate the role of tyrosine
kinases in the modulation of ICl,vol in cultured
astrocytes, we performed successive short (3-5 min) applications of
HOS in the presence or absence of different tyrosine kinase inhibitors
(with 2 mM ATP and 0.5 mM GTP in the pipette
solution). First, we established that repeated applications (up to
five) of HOS consistently activated ICl,vol with
no decrement in current amplitude (data not shown). Second, as
illustrated in Figure 5, we activated
ICl,vol with HOS in the absence or presence of
the tyrosine kinase inhibitor genistein (50 µM) and its
inactive analog, daidzein (50 µM), at concentrations
according to previous studies (for review, see Levitzki and Gazit,
1995). Figure 5A shows that, in the presence of daidzein,
HOS-activated ICl,vol was comparable to that
seen in absence of the drug, whereas concurrent application of HOS and
genistein not only prevented activation of
ICl,vol but also further decreased the control
current. The effects of genistein were reversible because HOS activated
ICl,vol within 7 min of genistein washout.
Figure 5B shows I/V relations for
ICl,vol measured from ramp commands in the
absence (Control) or presence of daidzein and
genistein and after drug washout. Genistein decreased the positive and
negative conductance of ICl,vol by 68 ± 14% (p = 0.015; n = 5) and by
69.9 ± 11.5% (p = 0.022; n = 5), respectively.
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Figure 5.
Tyrosine kinases are involved in the activation of
ICl,vol. A, Membrane current
evoked by four successive HOS stimulations (5 min in duration,
VH = 60 mV) recorded in the presence or in the absence of either daidzein (50 µM) or genistein (50 µM) in the external media [and (in mM) 63 Cl, 2 ATP, and 0.5 GTP in the pipette solution].
B, Mean I/V relations of
ICl,vol obtained in five cells before (),
during, and after (+) bath application of either daidzein (50 µM, ) or genistein (50 µM, ).
C, Mean I/V relations of
ICl,vol obtained in seven cells before
(), during, and after (+) bath application of either tyrphostin A1
(50 µM, ) or tyrphostin A23 (50 µM,
). Note that genistein and tyrphostin A23 strongly and reversibly
reduced ICl,vol, in contrast to their
inactive analogs, daidzein and tyrphostin A1, which had no significant
effect on ICl,vol.
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Similar experiments also were performed with another tyrosine kinase
inhibitor, tyrphostin A23 (50 µM), and its inactive
analog, tyrphostin A1 (50 µM), at concentrations
according to previous studies (for review, see Gazit et al., 1989)
(Fig. 5C). Tyrphostin A23 decreased the positive and
negative conductance of ICl,vol by 82.1 ± 6.3% (p = 0.001; n = 7) and by
76 ± 9.5% (p = 0.0026; n = 7), respectively. Interestingly, like genistein (see Fig. 5A), tyrphostin A23 acted very rapidly on
ICl,vol (within 3 min of preincubation), and,
like genistein, the effects of this drug were fully reversible. The
inhibitory action of genistein and tyrphostin A23 on
ICl,vol was specific to activation of a tyrosine kinase, because we did not obtain a significant change of
ICl,vol in the presence of the inactive analogs
of either genistein (daidzein) or tyrphostin A23 (tyrphostin A1).
Daidzein (50 µM) changed the positive and the negative
conductance of ICl,vol by 5.5 ± 10.5% (p = 0.41; n = 5) and by
5.4 ± 14.3% (p = 0.65;
n = 5), respectively, whereas tyrphostin A1 (50 µM) changed the positive and negative conductance of
ICl,vol by 7.3 ± 17.7%
(p = 0.27; n = 5) and by 12.0 ± 5.2% (p = 0.11;
n = 5), respectively.
Some tyrosine kinase signaling pathways can lead to MAP kinase
activation (Malarkey et al., 1995). Recent studies performed in
different cell types have shown that hypotonic stress activates MAP
kinases (Schliess et al., 1995, 1996; Noé et al., 1996; Tilly et
al., 1996). To test whether MAP kinase activation is integral to the
activation of ICl,vol, we exposed
astrocytes to successive short (3-5 min) applications of HOS in the
presence or in the absence of the MEK inhibitor PD 98059 (50 µM) at a concentration according to previous studies
(Alessi et al., 1995; Dudley et al., 1995). As shown in Figure
6A, 50 µM
PD 98059 strongly inhibited ICl,vol, and
continuous exposure (10-15 min) to PD 98059 prevented the activation
of HOS current. In contrast to the tyrosine kinase inhibitors, only a
partial recovery of ICl,vol was obtained after PD 98059 exposure. The slow effect and recovery from the actions of the
MEK inhibitor are consistent with previous reports of the pretreatment
time necessary for inhibitory actions (Alessi et al., 1995; Dudley et
al., 1995). Figure 6B shows I/V relations for the maximal current activated during two control applications of
HOS (in the absence of drug), during HOS stimulation after an initial
12-15 min exposure to PD 98059, and after washout of the MEK
inhibitor. Compared with controls, PD 98050 decreased the positive and
negative conductance of ICl,vol by 83.7 ± 6.2% (p = 0.005; n = 6) and by
88 ± 4.7% (p = 0.009; n = 6), respectively.
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Figure 6.
MAP kinase is involved in the activation of
ICl,vol. A, Membrane current
evoked by five successive HOS stimulations (3 min and 30 sec in
duration, VH = 60 mV) recorded in the
presence or in the absence of 50 µM PD 98059 in the
external media [and (in mM) 63 Cl, 2 ATP, and 0.5 GTP in the pipette solution]. B, Mean
I/V relations of ICl,vol
obtained in six cells before (Control 1, ;
Control 2, ), during (), and after (+) bath
application of PD 98059. Note that PD 98059 strongly inhibited
ICl,vol and that this inhibition was
partially reversible during wash.
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To characterize the link between astrocyte swelling and the activity of
MAP kinase further, we sought to determine whether treatments affecting
ICl,vol similarly affected MAP kinase
activation. To this end we used a gel shift assay to monitor the
activation of MAP kinases. Activation of Erk-1 and Erk-2 by
phosphorylation decreases their electrophoretic mobilities; this
"shift" can be visualized in Western blots because the activated
forms run at an increased apparent molecular weight. Figure
7A depicts the response of
Erk-1 and Erk-2 to hypo-osmolarity. In untreated, starved cultures (at
time 0), both Erk-1 and Erk-2 appeared as distinct bands with
approximate molecular weights of 44 and 42 kDa, respectively.
Hypo-osmotic solution induced an evident shift in mobility of both
Erk-1 and Erk-2 by 5 min, with maximal effect by 15-20 min. This is
denoted by the appearance of the upper bands, representing the
activated forms of these proteins. Under isotonic conditions there was
a very slight increase in activated (phosphorylated) Erk-1 and Erk-2.
This possibly resulted from the cellular stress associated with the
addition of new iso-osmotic medium (Schliess et al., 1996). We next
examined the role of tyrosine kinases and MEK on Erk activation by
hypo-osmotic treatment. The tyrosine kinase inhibitor genistein (50 µM) markedly depressed the mobility shift of Erk-1 and
Erk-2 when it was applied 15 min before and during stimulation with
hypo-osmotic solution (Fig. 7B), whereas the inactive
analog, daidzein, had no effect. Similarly, the MEK inhibitor PD 98059 (50 µM) also inhibited Erk-1 and Erk-2 activation in
hypo-osmotic solution (Fig. 7B).
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Figure 7.
Activation of MAP kinase in response to
hypo-osmotic conditions is sensitive to tyrosine kinase and MEK
inhibition. Astrocyte cultures were exposed to either iso-osmotic
conditions (Iso) or hypo-osmotic conditions
(Hypo) with or without other treatments for the time
periods indicated. Whole-cell extracts were separated by SDS-Page,
transferred to PVDF, and immunoblotted with either an anti-MAP kinase
antibody (A and B) or an
anti-phosphotyrosine antibody (C).
A indicates that the gel shift of p44 Erk-1 and p42
Erk-2 was greater in the Hypo condition, indicating
activation. In B and C, the cultures were
subjected to a number of different treatments as indicated: PD 98059 (PD, 50 µM), 30 min preincubation followed
by stimulation; DMSO 0.1%, 30 min preincubation followed by
stimulation; and daidzein (D, 50 µM) and
genistein (G, 50 µM), 15 min preincubation
followed by stimulation. The decrease in the intensity of the
upper bands representing activated Erk-1 and Erk-2
(B) indicates that MAP kinase activity was
attenuated in these treatments. In C, the increased
tyrosine phosphorylation of Erk-2 in hypo-osmotic solution is apparent
at 20 min. PD 98059 and genistein dramatically reduced tyrosine
phosphorylation of Erk-2. Other apparently immunoreactive proteins were
not affected by these treatments (C).
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Because tyrosine phosphorylation of MAP kinase is a prerequisite for
activity, the presence of enhanced tyrosine phosphorylation of the Erks
has been used as an indicator of kinase activation. Anti-phosphotyrosine Western blotting confirmed the activation of Erk-2
in response to hypo-osmotic conditions, as well as the inhibitory
effect of genistein and PD 98059 (Fig. 7C). The Western blots of Erk-1 were too faint to give a clear indication of activity changes.
| |
DISCUSSION |
The present report describes the second messenger systems
modulating an outwardly rectifying Cl current
activated by hypo-osmotic solution in cultured astrocytes. Current
activation was correlated with phosphorylation processes but did not
require changes in intracellular Ca2+. This study
demonstrates that activation of tyrosine and MAP kinases plays a
crucial role in the activation of the swelling-activated Cl current. Western immunoblots confirmed that MAP
kinases were activated by swelling and showed that activation of MAP
kinases depended on the activation of tyrosine kinases and MEK. This
study provides the first demonstration that MAP kinases and tyrosine kinases are involved in the activation of the volume-activated Cl current in astrocytes.
Properties of ICl,vol in
cultured astrocytes
Activation of ion channels appears to be a primary step in the
regulatory volume decrease (RVD) that follows an increase in cell
volume. In many cells RVD requires coactivation of
K+ and Cl channels and a net
efflux of KCl (Roy and Banderali, 1994) (for review, see Nilius et al.,
1996; Strange et al., 1996). Activation of volume-activated
Cl channels has been described in a variety of
cell types in response to challenge with extracellular hypotonic
solution (for review, see Nilius et al., 1996; Strange et al., 1996),
increased pressure applied to the cell membrane (Hagiwara et al.,
1992), or morphological changes (Lascola and Kraig, 1996). The
characteristics of the volume-activated Cl current
vary among cells, but, in general, the current is outwardly rectifying
and does not appear to be mediated by intracellular Ca2+ (for review, see Nilius et al., 1996; Strange
et al., 1996). Volume-activated anion currents have been described in
astrocytes cultures, astrocytomas, and glioma cells (Pasantes-Morales
et al., 1994; Bakhramov et al., 1995; Strange and Jackson, 1995; Lascola and Kraig, 1996). In the present study we characterized the
Cl current activated by swelling with HOS in
cultured cortical astrocytes. All experiments were performed in the
absence of Na+ and K+ in the
internal and external media and in the presence of
K+ channel blockers in the internal
(Cs+) and the external media (TEA and
Ba2+). We showed that bath application of HOS
solution activated an outwardly rectifying current, which was blocked
by Cl channel inhibitors (NPPB and SITS) and
displayed a reversal potential close to the predicted chloride
equilibrium potential calculated with the Nernst equation. Current
activation was independent of intracellular [Ca2+]
changes, because its activation also occurred in the presence of a
Ca2+ chelator (BAPTA) in the internal pipette
solution.
Therefore, the Cl current activated by
hypo-osmotic-induced swelling in cultured astrocytes has similar
features to Cl currents previously described in
astrocytes and other cell types (Lascola and Kraig, 1996) (for review,
see Nilius et al., 1996; Strange et al., 1996).
The activation of ICl,vol involves
phosphorylation processes and the MAP kinases signaling pathway
The stable activation of volume-sensitive Cl
currents in many, but not all, cell types requires intracellular ATP
(but see Kelly et al., 1994; Leaney et al., 1997). For example, in
cardiac and cortical-collecting duct cells, the hydrolyzable form of
ATP is required (Sorota, 1995; Meyer and Korbmacher, 1996), but in other cells types nonhydrolyzable ATP analogs can substitute for ATP
(Jackson et al., 1994; Oike et al., 1994; Oiki et al., 1994). It has
been suggested that ATP is linked to a nonhydrolytic binding site (for
review, see Strange et al., 1996). To clarify the role of ATP in the
activation of ICl,vol in cultured astrocytes, we performed a set of experiments either in the presence or in the absence
of ATP or in the presence of the nonhydrolyzable ATP analog, AMP-PNP.
We report that, in cultured astrocytes, ICl,vol
activation depends on the hydrolyzable form of ATP in the internal
solution, because a stable activation of ICl,vol
was obtained in the presence of ATP, but not in the absence of ATP or
in the presence of AMP-PNP, a nonhydrolyzable ATP analog. This clearly
suggests that in cultured astrocytes the activation of
ICl,vol involved phosphorylation-dependent processes.
The role of phosphorylation processes in the activation of the
volume-activated Cl current has been studied
extensively. Although it is well established in many cells that PKC and
PKA signaling pathways are not involved in the activation of this
current (Hagiwara et al., 1992; Kelly et al., 1994; Nilius et al.,
1994; Gosling et al., 1995; Szucs et al., 1996; Leaney et al., 1997)
(but see Schwiebert et al., 1994; Kinard and Satin, 1995; Verdon et
al., 1995), they may be involved in its modulation (McCann et al.,
1989; Hardy et al., 1994; Du and Sorota, 1997). The role of tyrosine
kinase signaling pathways also has been tested, and it was shown that
tyrosine kinase activation is a key step in the activation of the
volume-sensitive Cl conductance in Intestine 407 and cardiac cells (Tilly et al., 1993; Sorota, 1995), but not in other
cell types (Gosling et al., 1995; Szucs et al., 1996). In astrocytes
the role of tyrosine kinases in the activation of
ICl,vol has not been studied. However, because
biochemical studies have demonstrated clearly that MAP kinase is
activated by swelling (Tilly et al., 1993; Schliess et al., 1995, 1996;
Noé et al., 1996) and because MAP kinases are linked to some
tyrosine kinase signaling pathways (for review, see Graves et al.,
1995), we hypothesized that both tyrosine and MAP kinase activation are
necessary for the activation of ICl,vol in
astrocytes.
In the present study we show that, as in cardiac cells (Sorota, 1995),
ICl,vol activation in astrocytes depends on the
activation of tyrosine kinases. In the presence of specific inhibitors
of tyrosine kinases (genistein and tyrphostin A23),
ICl,vol was reversibly depressed while the
respective inactive structural analogs of these compounds, daidzein and
tyrphostin A1 (Negrescu et al., 1995), were ineffective. Furthermore,
we demonstrate that MAP kinases also are involved in the activation of
ICl,vol, because the specific MEK
inhibitor PD 98059 (Dudley et al., 1995) reversibly inhibited the
current. Finally, we confirmed, using Western immunoblots, that MAP
kinase phosphorylation was induced by the hypo-osmotic solution and
that the activation of MAP kinases depended on tyrosine kinase and MEK
activity, because it was blocked by the tyrosine kinase inhibitor
genistein (and not by its structural inactive analog daidzein) and a
specific inhibitor of MEK, PD 98059. These results point as well to the
relationship of tyrosine kinases to MAP kinases. The tyrosine kinase
inhibitors used in this study do not act directly on MEK (Cox et al.,
1996); however, they inhibited the activation of MAP kinase, suggesting
that they act upstream of MEK.
Several studies have shown that the swelling of astrocytes also induces
changes in the cytoskeleton. Recent studies indicate that
reorganization of the F-actin cytoskeleton may play an important role
in the activation of ICl,vol (Schwiebert et al.,
1994; Lascola and Kraig, 1996; Tilly et al., 1996), possibly via the
regulatory protein plCln (Krapivinsky et al., 1994). It
also has been shown that the p21rho signaling
cascade, implicated in tyrosine kinase-mediated F-actin reorganization,
is involved in the activation of ICl,vol
(Lascola and Kraig, 1996). Although it is premature to speculate that
the dependence of ICl,vol on tyrosine kinases
may be linked to these events, the cytoskeleton does present an
attractive future route for further study into signaling and
ICl,vol.
Concluding remarks
The Cl permeability triggered by osmotic
swelling is a requisite for RVD in astrocytes (Pasantes-Morales et al.,
1994) as well as in other cell types (Nilius et al., 1996; Strange et
al., 1996). Delineating the signaling pathways in the activation of ICl,vol may be important in understanding the
mechanisms involved in RVD. In the present study we demonstrate that
both tyrosine and MAP kinases play a crucial role in the activation of
ICl,vol, and we show for the first time
that the MAP kinase cascade is involved in the activation of an ionic
channel. We also demonstrate that tyrosine kinases are likely upstream
in the signaling pathway of the MAP kinase-dependent activation of
ICl,vol in astrocytes. Further investigations
will be required to determine the specific target of the tyrosine and
MAP kinases involved in the activation of
ICl,vol, as well as the sensor leading to
the activation of these second messenger cascades during the cell
swelling.
| |
FOOTNOTES |
Received Aug. 21, 1997; revised Nov. 19, 1997; accepted Nov. 20, 1997.
This work was supported financially by the Alberta Heritage Foundation
for Medical Research (AHFMR), the Institut National de la Santé
et de la Recherche Médicale (INSERM), and the Medical Research
Council of Canada (MRC). B.A.M. is an MRC Senior Scientist and an AHFMR
Scientist. M.E.M.K. was an AHFMR visiting scientist. We thank D. Feighan for technical assistance and for preparing astrocyte cultures.
We also thank Drs. S. Williams and M. Hollenberg for helpful
discussions.
Correspondence should be addressed to Dr. Brian A. MacVicar at the
above address.
| |
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Top
Abstract
Introduction
