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The Journal of Neuroscience, March 1, 1998, 18(5):1679-1692
Cytoskeletal Actin Gates a Cl
Channel in
Neocortical Astrocytes
Christopher D.
Lascola1,
Deborah J.
Nelson1, 2, and
Richard P.
Kraig1, 2
1 Committee on Neurobiology, and
2 Departments of Neurology and Pharmacological and
Physiological Sciences, The University of Chicago, Chicago, Illinois
60637
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ABSTRACT |
Increases in astroglial Cl
conductance
accompany changes in cell morphology and disassembly of cytoskeletal
actin, but Cl channels underlying these
conductance increases have not been described. We characterize an
outwardly rectifying Cl channel in rodent
neocortical cultured astrocytes and describe how cell shape and
cytoskeletal actin modulate channel gating. In inside-out patch-clamp
recordings from cultured astrocytes, outwardly rectifying
Cl channels either were spontaneously active or
inducible in quiescent patches by depolarizing voltage steps. Average
single-channel conductance was 36 pS between 
60 and 80 mV and was
75 pS between 60 and 80 mV in symmetrical (150 mM NaCl)
solutions. The permeability ratio
(PNa/PCl)
was 0.14 at lower ionic strength but increased at higher salt
concentrations. Both ATP and 4,4-diisothiocyanostilbene-2,2'-disulfonic acid produced a flicker block, whereas Zn2+ produced
complete inhibition of channel activity.
The frequency of observing both spontaneous and inducible
Cl channel activity was markedly higher in
stellate than in flat, polygonally shaped astrocytes. In addition,
cytoskeletal actin modulated channel open-state probability
(PO) and conductance at negative membrane
potentials, controlling the degree of outward rectification. Direct
application of phalloidin, which stabilizes actin, preserved low
PO and promoted lower conductance levels at
negative potentials. Lower PO also was induced
by direct application of polymerized actin. The actions of phalloidin
and actin were reversed by coapplication of gelsolin and cytochalasin
D, respectively. These results provide the first report of an outwardly
rectifying Cl channel in neocortical astrocytes
and demonstrate how changes in cell shape and cytoskeletal actin may
control Cl conductance in these cells.
Key words:
astrocytes; chloride channels; actin; cytoskeleton; phalloidin; gelsolin patch clamp
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INTRODUCTION |
Changes in Cl
conductance are likely to have profound consequences for astroglial
function in brain. Cl serves as an important
counter-ion to K+ movement and is a transported
species on ion exchangers and transporters carrying
H+ equivalents (Kimelberg et al., 1986
). Because
astrocytes are believed to regulate neuronal activity via active
maintenance of interstitial K+ and pH (Orkand et
al., 1966; Chesler, 1990), changes in Cl
conductance may be an important means by which astrocytes shape brain
cell signaling.
Previously, we have shown that increases in Cl
conductance accompany changes in astroglial morphology and the
disruption of the actin cytoskeleton (Lascola and Kraig, 1996). These
results suggested that cytoskeletal actin may be part of a novel
signaling mechanism in astrocytes, and the results added to the growing concept that cytoskeletal elements can modulate ion channel function (Cantiello et al., 1991; Johnson and Byerly, 1993; Rosenmund and Westbrook, 1993; Suzuki et al., 1993; Haussler et al., 1994; Schwiebert et al., 1994; Levitan et al., 1995; Ismailov et al., 1997). A dynamic
cytoskeleton therefore may be one mechanism by which astrocytes couple
changes in cell and brain function to the dramatic changes in their
morphology accompanying such processes as reactive astrocytosis.
Cl channels that may be responsible for
morphology-dependent Cl conductance increases in
astrocytes have not yet been characterized at the single-channel level.
To date, three Cl channels have been reported in
cortical astrocytes, but the activation and kinetic properties of these
channels do not match the characteristics of morphology-dependent and
actin-dependent whole-cell Cl currents. Bormann
and Kettenmann (1988) have analyzed single Cl
channels activated by GABA in astrocytes that possess kinetic behavior
and conductance substates similar to those described for neuronal
GABA-activated Cl channels. Sonnhof (1987), Nowak
et al. (1987), and Jalonen (1993) have studied a large conductance
astroglial anion channel found in many cell types, the open-state
probability of which rapidly declines with polarization to either
positive or negative potentials. Finally, Nowak et al. (1987) reported
the activation of a small, 5 pS channel in outside-out patches from
cultured rat astrocytes that gated at hyperpolarizing potentials.
Morphology-dependent whole-cell Cl currents in
astrocytes have either a linear or outwardly rectifying current-voltage relationship, variably express voltage-dependent activation and inactivation kinetics, and do not require GABA (Lascola
and Kraig, 1996).
In this paper we characterize an outwardly rectifying
Cl-selective channel (ORCC), previously
undescribed in astrocytes, that shares kinetic and pharmacological
features with morphology-dependent and actin-dependent astroglial
whole-cell Cl currents. Moreover, the frequency of
observing channel activity increases in patches excised from
morphologically transformed cells. In addition, we also show that
alteration of actin polymerization in membrane patches controls channel
rectification by modulating channel PO and
conductance, particularly at negative membrane potentials. Because
astrocytes are nonexcitable cells within the nervous system and
therefore function at membrane potentials exclusively within a negative
voltage range, the results described below suggest that cytoskeletal
actin may gate ~80-90% of whole-cell Cl
current caused by ORCCs at membrane potentials physiologically and
pathologically relevant to astrocytic function.
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MATERIALS AND METHODS |
Cell culture preparation. Neocortical type 1 astrocytes were prepared according to the methods of McCarthy and
DeVellis (1980), as described previously (Lascola and Kraig, 1996).
Recordings were made between days 3 and 7 after secondary plating and
before the cells reached monolayer confluence. Both flat, polygonal
(control) and stellate astrocytes were examined for
Cl channels. Control cells were introduced
directly to recording solutions from DMEM. Control cells were
transformed into a stellate morphology by incubation in Ringer's
solution for 2-6 hr (Moonen, 1975; Lascola and Kraig, 1996) at 37°C.
Stellate cells had rounded-up cell bodies and multiple cell
processes.
Solutions. For most inside-out patch-clamp recordings, the
following solutions were used. Bath contained (in mM): 150 NaCl, 2 MgCl2, and 10 N-tris(hydroxymethyl) methyl-2-aminoethane-sulfonic acid
(TES), pH 7.4. Pipette contained (in mM): 150 NaCl, 2 CaCl2, 10 TES, and 0.1 GdCl3. The choice
of TES instead of HEPES as a buffer was based on preliminary
experiments demonstrating that HEPES reduced the outward rectifier
conductance by ~25-40%, corroborating the results of Hanrahan and
Tabcharani (1990), who showed significant inhibition of an epithelial
ORCC by HEPES, but not by TES. Gd3+, a general
cation channel blocker, was included in the pipette to block cation and
stretch-activated channels (Yang and Sachs, 1989) and to help stabilize
the membrane-pipette seal. Gd3+ did not
significantly change the open or closed dwell times, burst durations,
or open-state probability of the astroglial ORCC.
To examine Cl selectivity in inside-out patch
experiments, we changed Cl concentrations by
isosmotic replacement of NaCl in either the bath or pipette solutions.
Patch integrity was stable during solution changes to higher ionic
strengths (e.g., 300-600 mM [NaCl]o). Patch instability accompanied the bath NaCl exchange from 150 to 50 mM; therefore, in these experiments 200 mM
sucrose was added to the 50 mM NaCl solution to maintain
osmolarity.
Electrophysiology. Cl channel currents
were recorded by using the inside-out and cell-attached patch-clamp
techniques (Hamill et al., 1981). Pipettes were pulled in four to five
stages, heat-polished on a Narishige microforge (model MF-83,
Setagaya-KU, Tokyo, Japan), and had resistances of 3-6 M
when
filled with a 150 mM NaCl solution. The bath was grounded
via an agar bridge having the same NaCl composition as the pipette
solution. Membrane potential was set manually on the Axopatch 200A
(Axon Instruments, Foster City, CA), and neither series resistance nor
capacitance compensation was used.
Electrophysiological recordings were performed at room temperature
(20-22°C). Currents were recorded directly onto hard disk or optical
disk via a Digidata 1200 A/D converter (Axon Instruments) interfaced
with an IBM-compatible computer (AST Premmia 486/66d; Irvine, CA).
Single-channel currents were low-pass-filtered with an eight-pole
Bessel filter (902LPF, Frequency Devices, Haverhill, MA) set at a
corner frequency (3 dB) of 300-600 Hz and sampled at 100 µsec per
point. Data were analyzed on a 486/66 computer (AST), using the pCLAMP
(Axon Instruments) software suite and ORIGIN statistical software
(Microcal Software, Northhampton, MA). Single-channel openings were
evaluated by using a threshold method in FETCHAN (pCLAMP) set at 50%
of the single-channel open current. A relative measure of time spent in
the open state (PO) was calculated
quantitatively in pSTAT (pCLAMP) from event list files generated in
FETCHAN.
The relative permeability of the channel for Cl
over Na+
(PCl/PNa)
was calculated from zero current potentials
(VO) by using the Goldman-Hodgkin-Katz
equation (Goldman, 1943). Single-channel current-voltage
(I-V) plots were fit by using a third-order
polynomial, and zero current potentials were determined from the
abscissa intersections of the polynomials. Cl and
Na+ activities were calculated by the modified
Debye-Huckel formula (Robinson and Stokes, 1965). Slope conductances
were determined by linear regression over the voltage ranges of
interest.
Open and closed time distributions were fit with least-squares fitting
routines (Simplex and Levenberg-Marquardt) in pSTAT and ORIGIN. Curve
fitting was performed first in pSTAT (Axon Instruments), and then it
was performed again in ORIGIN (Microcal); the time and proportionality
constants obtained from both fitting algorithms were compared for
agreement.
Open, closed, and burst duration histograms were fit according to the
methods of Colquhoun and Hawkes (1981). Determination of the interburst
interval, the minimum value of a closed duration that separates bursts,
was estimated according to the methods of Sigurdson et al. (1987) and
was performed in pSTAT.
Statistics. Data were expressed as mean ± SEM.
Student's t test was used to evaluate significance.
Materials. Gelsolin (Sigma, St. Louis, MO) was used at a
final activity of 1 U/ml. Gelsolin was made up in a solution of 150 NaCl, 20 mM Tris, 200 µM
CaCl2, 200 µM MgATP, and 0.2 mM EGTA (final free [Ca2+] = 45 µM), pH 7.4 (NaOH), and frozen in 5 µl (5 µg/ml
protein) aliquots at 70°C until the time of experiment. Gelsolin
was introduced directly to the 500 µl recording bath.
Lyophilized actin (Sigma) was dissolved in a low ionic strength buffer
(Cantiello et al., 1991) containing (in mM) 0.5 Tris-Cl, 0.2 CaCl2, 0.5 ATP, and 0.5 
-mercaptoethanol, pH
8.0, at a concentration of 50 mg/ml in 10 µl aliquots and rapidly was
frozen to 70°C. To make a mixed solution of long actin polymers,
short polymers, and monomers, we diluted aliquots into a 100 µl
solution containing 150 NaCl, 2 Mg ATP, and 10 TES, pH 7.4 (NaOH), and
incubated the solution for 12 hr at 37°C. The mixed polymer/monomer
solution was added to the 500 µl recording bath for a final
concentration of 1 mg/ml.
Phalloidin and cytochalasin D (both from Sigma) were solubilized in
methanol and DMSO, respectively, and stored in stock solutions at
20°C. Fetal calf serum for culture media was obtained from Life
Technologies (Grand Island, NY).
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RESULTS |
Activation in inside-out patches
When inside-out patches were excised from cultured astrocytes,
ORCCs either were spontaneously active or were activated by applying
depolarizing voltage pulses to 
60 mV lasting seconds to minutes. Once
activated, ORCCs remained active and quickly progressed toward a
relatively high open-state probability at both positive and negative
membrane potentials (Fig.
1A,B). This pattern of
either spontaneous activity or activation after depolarizing voltage
pulses is characteristic of ORCCs that are observed in epithelial
(Tabcharani and Hanrahan, 1991), neuronal (Franciolini and Nonner,
1987, 1994), and muscle cells (Fahlke et al., 1992).
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Figure 1.
Outwardly rectifying Cl
channel (ORCC) activity in inside-out patches from cultured rat
astrocytes. ORCCs either were spontaneously active or required
high-voltage pulses for activation (see Results) when patches were
excised in symmetrical 150 mM NaCl solutions. A, Noncontiguous current traces of the ORCC at 1 sec
intervals at membrane potentials from 60 to 60 mV in 20 mV
increments. The channel from which these traces were obtained was
activated after the patch potential was stepped to 60 mV for 45 sec.
The C adjacent to the dotted line
indicates the closed state of the channel. B, Current
traces of 10 sec from which the 40 and 40 mV traces were taken in
A are displayed to illustrate channel activity over an
extended period of time. C, Single-channel
I-V plot demonstrating the outward rectification of the
channel. Slope conductance was 36 pS between 80 and 60 mV and was
75 pS between 60 and 80 mV.
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To provide a basis with which to compare channel expression and
activity in patches excised from different experimental populations of
astrocytes, we used a standardized voltage protocol similar to that
developed by Welsh et al. (1989) and Tabcharani and Hanrahan (1991) for
epithelial cells. After patch excision, patches were held at 30 mV
for 2 min. If channel activity was observed, channels were classified
as "spontaneous." The time period of 2 min was established by
experiments demonstrating that channels not active within 2 min at 30
mV thereafter would never become "spontaneously" active. Patches
without spontaneously active channels were stepped to 60 mV next and
held at this potential for 1 min. If no channel activity was observed
under these conditions, then the patch was stepped back and forth
between 0 and 60 mV for 10 sec at each potential for an additional 1 min. Stepping back to 0 mV was necessary because in ~20-30% of
patches Cl channels exhibited
depolarization-dependent inactivation (Fig. 2) at potentials 60 mV. If channel
activity still was not observed after these steps, the same pattern of
voltage pulses was repeated to 90 mV and then to 120 mV. Channels in
patches pulsed to 60 mV were classified as "voltage-activated."
Patches not demonstrating channel activity after voltage pulses to 120 mV were assumed not to contain Cl channels.
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Figure 2.
Depolarization-dependent inactivation of the ORCC.
A, Amplitude histograms of current traces at 40 and 40 mV that were shown in Figure 1B. The histograms
were fit with second-order gaussians. Mean current at 40 mV was 2.9 pA
and at 40 mV was 1.75 pA, indicating a single conductance level of 72 and 44 pS at 40 and 40 mV, respectively. B,
Depolarization-induced inactivation is shown in a graph of PO versus membrane potential (mV) for the
channel shown in Figure 1. PO within the
first second (instantaneous PO)
versus mV is shown by the filled circles. Steady-state
availability is indicated by the filled squares.
C, Current records taken from the channel in Figure 1
show the channel first being stepped from 0 to 80 mV and then from 80 to 30 mV. The voltage protocol is shown above the
current traces. At 80 mV (the top current trace), the
channel at first has high open-state probability
(PO) before inactivating <1 sec
after the voltage step has been made. The channel thereafter remained
inactivated for 30 sec (the remainder of the record is not displayed).
Then the patch was stepped from 80 to 30 mV and the channel was
reopened, maintaining a high PO.
D, Whole-cell Cl currents in
astrocytes transformed in warm Ringer's solution (Lascola and Kraig,
1996) into a stellate morphology also demonstrate voltage-dependent
inactivation at positive potentials.
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In most inside-out patches showing Cl channel
activity, only one channel was observed. Occasionally, inside-out
patches possessed three or more ORCCs. Because of this and the
observation that some channels apparently would enter long closed
states of several tens of seconds to several minutes before
reactivation, the actual number of channels per patch could not be
determined unequivocally.
Under the conditions used in these experiments (i.e.,
Ca2+-free and K+-free solutions),
excised inside-out patches from astrocytes demonstrated little channel
activity other than the ORCC. On occasion, we observed a large
conductance "maxi" anion channel previously described by several
other investigators (Nowak et al., 1987; Sonnhof, 1987; Jalonen, 1993).
This channel had a conductance with an order of magnitude larger than
the ORCC and rapidly inactivated when the membrane potential was
stepped more than ±20 mV away from 0 mV. Although this channel was
readily distinguishable from the ORCC, membrane patches containing
these channels were discarded.
Conductance and voltage-dependent inactivation
Figure 1A demonstrates 1-sec-long current traces
of the ORCC at membrane potentials from 60 to 60 mV (in 20 mV
increments). The channel from which these traces were obtained was
activated after the patch potential was stepped to 60 mV for 45 sec. At this resolution it is evident that the channel spends most of its time
in the open state and that there are both slow and fast closures that
occur relatively infrequently. Current traces induced during voltage
steps of opposite polarity are displayed alongside each other to
emphasize the outward rectification of the channel. Even at ± 20 mV the larger conductance at the positive potential is readily
discernible.
Figure 1B demonstrates two 10 sec traces at +40 and
40 mV from which the ±40 mV traces in Figure 1A
were taken. Visual inspection of these traces provides a first
indication that the current through the ORCC is modulated by a gating
process with complex kinetics. Openings occurred in bursts lasting from
several tens of milliseconds to several seconds, and the bursts
contained many short closures. Figure 1C is an
I-V plot of the channel depicted in Figure 1, A
and B. The slope conductance at negative potentials between 60 and 80 mV was 36 pS in symmetrical 150 NaCl solutions and 75 pS
at positive potentials between 60 and 80 mV.
Figure 2A shows representative amplitude histograms
of channel activity during the 40 and 40 mV traces shown in Figure
1B. The histograms were fit with second-order
gaussians. Both plots demonstrate two clearly delineated peaks. The
peak at 0 pA in each graph represents closed events, and the other peak
represents open events. The current at 40 mV was 2.9 pA, and that at
40 mV was 1.75 pA, indicating conductance levels of 72 and 44 pS at
40 and 40 mV, respectively. Mean conductance in 26 patches at 40 mV
was 45 ± 1.0 pS (range, 35-53 pS) and 64 ± 1.2 pS (range, 54-74 pS) at 40 mV in symmetrical 150 mM NaCl solutions,
with 10 mM TES as a pH buffer.
The amplitude histograms in Figure 2A provide an
indication of the relative PO of the channel at
positive and negative potentials. At 40 mV, the channel spent most of
its time in the open state, corresponding to a
PO of 0.92 for a 30 sec recording at this
potential. Mean PO for 26 patches at 40 mV was
0.81 ± 0.04 (range, 0.43-0.97). At 40 mV, a reduced
PO of 0.75 was observed. The mean
PO for 26 patches at 40 mV was 0.70 ± 0.04 (range, 0.25-0.91). The trend over the course of several
recordings showed an increase in PO from
negative to positive potentials.
At positive potentials, the reduction in PO is
attributable in part to the presence of depolarization-dependent
inactivation. Depolarization-dependent inactivation was observed in
~20-30% of patches (Fig. 2C), greatly reducing
open-state probability (PO) at high
positive potentials. The range of percentages is approximate, because
this inactivation process was highly variable both in the rate of
inactivation and the voltage dependence of inactivation, and expression
was also variable. If present in inside-out patches,
depolarization-dependent inactivation typically was first evident at 60 mV. As patch potential was clamped to increasingly positive membrane
potentials, inactivation became progressively faster. In Figure
2C, current records are shown in which the patch potential
is stepped first from 0 to 80 mV and then from 80 to 30 mV. At 80 mV
(the top current trace), the channel first exhibited a high
open-state probability before inactivating <1 sec after the voltage
step was made. The channel thereafter remained inactivated as the patch
potential was held at 80 mV. Then the patch was stepped down to 30 mV.
At this potential the channel reopened, maintained a high open-state
probability, and did not inactivate. Typically, ORCCs had only to be
stepped back to membrane potentials just outside the range at which
depolarization-dependent inactivation was observed (i.e., 
60 mV) to
remove inactivation.
An example of time-dependent and voltage-dependent inactivation in a
whole-cell recording from a stellate astrocyte is demonstrated in
Figure 2D. Depolarization-dependent inactivation in
whole-cell recordings showed considerable variability and varied not
only across cells but also within recordings (Lascola and Kraig, 1996), paralleling the considerable variability also observed in single Cl channels from excised patches.
Cl selectivity
Figure 3A represents an
experiment in which an inside-out patch was exposed first to
symmetrical 150 mM NaCl solutions, and then the bath
solution was switched to one in which 100 mM NaCl was
replaced iso-osmotically with sucrose. The reversal potential shift for
three patches under these conditions was 20.1 ± 0.7 mV, showing
a PNa/PCl of
~0.14. The predicted shift for a perfectly Cl
selective conductance under these conditions is 28 mV.
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Figure 3.
Cl selectivity of the ORCC.
A, The selectivity of the ORCC was assessed by varying
the internal concentration of Na and Cl ions. Initially, the patch was
bathed in symmetrical 150 mM NaCl solutions. The
I-V relationship in these solutions reverses at 0 mV
and is indicated by the filled circles. Then the bath
was switched to one containing only 50 mM NaCl
(unfilled circles) while the patch solution was held
constant. The reversal potential for three patches that followed this
solution change was 20 ± 0.7 mV (predicted NernstCl = 28 mV), suggesting a
PNa/PCl of ~0.14. B, I-V relationships
demonstrating shifts in reversal potential after bath solution changes
from 300 to 600 mM NaCl (unfilled circles)
and from 300 to 150 mM NaCl (unfilled
squares) while holding the pipette constant at 300 mM NaCl. The I-V relationship when 300 mM NaCl was present in both the bath and pipette is shown by the filled circles. When the bath was switched first
to 600 mM NaCl, the zero current potential shifted to
11.2 ± 0.8 mV (n = 4; predicted
NernstCl = 17 mV), indicating a
PNa/PCl of
0.18. After the bath was switched to 150 NaCl, the mean reversal was 12.0 ± 0.6 mV (n = 4; predicted
NernstCl = 17 mV), and the
PNa/PCl was ~0.2. C, Comparison of the I-V
relationship in 150 mM NaCl solutions (filled
circles) with one obtained in symmetrical 150 NMDG-Cl
solutions (n = 12; unfilled
circles). In the Na-free solutions, note the decrease in both
inward and outward current amplitude, although the
Cl concentrations remain the same.
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The finite cation permeability increased in solutions of increasing
ionic strength. Figure 3B details the results of a
representative experiment in which a patch was exposed first to
symmetrical 300 mM NaCl solutions; then the bath was
shifted sequentially to one with 150 NaCl and finally to one with 600 NaCl. The pipette solution was fixed at 300 NaCl. After the first
solution change to 150 NaCl, the zero current potential shifted to
12.0 ± 0.6 mV (n = 4; predicted Nernst = 17 mV). After the second solution change, the zero current potential
shifted to 11.2 ± 0.8 (n = 4; predicted Nernst = 17 mV). The
PNa/PCl from the
first shift to 150 NaCl was 0.18; the
PNa/PCl for the
second shift was 0.22. Thus, the permeability of the astroglial outward
rectifier to Na+ seemed to increase with
progressively higher salt concentrations.
Both inward and outward currents (Cl current from
the pipette to the bath) were reduced when the concentration of NaCl in the bath was lower than that in the pipette, as illustrated in Figure
3, A and B. Interestingly, reduction in
inward current amplitude after an increase in bath NaCl with
respect to pipette NaCl was not observed, as demonstrated in Figure 3.
This paradoxical reduction in both inward and outward current in
asymmetrical solutions has been reported in selectivity experiments for
outwardly rectifying Cl channels in epithelial
cells (Halm et al., 1988; Tabcharani et al., 1989).
Channel inhibition by DIDS, ATP, and Zn2+
Figure 4 demonstrates block of the
outward rectifier by three agents known to inhibit single-channel and
whole-cell anion currents in other preparations, including
morphology-dependent whole-cell Cl currents
(Lascola and Kraig, 1996). Figure 4A shows
representative current traces from a patch in which two channels are
active at 30 mV and only one is active at 30 mV. One minute after the
application of 200 µM DIDS to the "intracellular"
face of the patch, the current traces showed a rapid closure or
flickering that was present at both positive and negative potentials.
The flickering block persisted, after washout of submaximal inhibitory
concentrations of DIDS, indicating the irreversibility of the block
(n = 3). In Figure 4B, current traces
are shown at 40 and 40 mV before and after the addition of 2 mM ATP to the bath. Submaximal inhibitory concentrations of
ATP produced a flickering channel block similar to that observed with
DIDS. ATP block was completely reversible after washout of the
nucleotide (data not shown) (n = 3). Block with ATP
from the intracellular face of the membrane suggests that intracellular levels of ATP could have a direct role in modulating
Cl permeability.
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Figure 4.
Block by DIDS, ATP, and Zn2+.
A, Channel inhibition of 200 µM DIDS 1 min
after application to the intracellular face of the patch
(n = 3). DIDS produced a flickering block that was
voltage-independent and irreversible. B, Channel
inhibition by 2 mM ATP immediately after its addition to
the intracellular/bath solution (n = 3). ATP
produced an increase in open channel noise, especially at negative
potentials. Some flicker block (rapid, complete closures) also was
evident. Block was also voltage-independent but, in this case,
completely reversible after washout of the nucleotide.
C, The addition of 1 mM
Zn2+ completely abolished single-channel currents in
<1 min (n = 3). This effect was reversible, but
only after several minutes of washout. The C adjacent to
the dotted line indicates the closed state of the
channel.
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Figure 4C shows channel inhibition by
Zn2+. The addition of 1 mM
Zn2+ to the bath completely abolished the
single-channel currents in <1 min (n = 3). This effect
was reversible but only after several minutes of solution exchange.
Morphology-dependent expression
Inside-out patches from both polygonal and stellate astrocytes
were examined for ORCC activity. Once activated, ORCCs from polygonal
and stellate cells did not demonstrate any differences in outwardly
rectifying voltage-dependent channel properties. Cells of each
morphology, however, did exhibit statistically significant differences
in Cl channel expression. Figure
5 illustrates the difference in
expression of both spontaneously active and voltage-activated ORCCs in
excised inside-out patches from polygonal and stellate astrocytes.
Outwardly rectifying channels that were spontaneously active
immediately after patch excision were compared with the expression of
channels requiring voltage activation. Spontaneously active
Cl channels were observed in 5% of polygonal
cells (n = 59), as compared with 19% of stellate cells
(n = 32). This difference in expression may reflect
either an increase in the activity of the physiological mechanism or
mechanisms responsible for activating these channels in stellate cells
or an increase in the number of channels per patch. A comparison of the
percentage of expression of voltage-activated channels provides some
insight into these possibilities. Although the voltage activation
protocol was standardized for both morphological classes of cells (see
above), channel expression was still significantly greater in stellate
cells (81%), as compared with polygonal cells (58%). This suggests
that the actual number of channels per excised patch membrane may be
greater from stellate cells than from flat cells.
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Figure 5.
Morphology-dependent expression of the ORCC. Shown
is the percentage of expression of both spontaneous and induced Cl
channel activity in excised and cell-attached patches, which differed between polygonal cells and stellate cells. Percentages represent the
cumulative proportion of patches expressing Cl channels. In excised
inside-out recordings, spontaneously active channels were more than
three times as likely to be observed in patches excised from stellate
cells (19%) than from polygonal cells (5%) (n = 91). When a standardized voltage activation protocol was used, depolarization-induced Cl channel activity was also greater in patches
from stellate cells (81%), as compared with polygonal cells
(58%).
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Spontaneous low-to-high PO transitions at
negative potentials
After patch excision, spontaneous or depolarization-induced ORCCs
usually exhibited stationary behavior. On occasion, however, ORCCs
sometimes were observed at first to enter a state in which PO was dramatically lower at negative potentials
but only slightly reduced at positive potentials (Fig.
6). The low PO
state just after channel activation, seen especially at negative
potentials, was observed in approximately one-third of inside-out
patches and was highly variable both in PO and
in duration. Sometimes, it was observed briefly at a single negative
potential within the first few seconds of recording, before channel
activity became relatively fixed at a higher PO.
In other patches a state of low PO could be
observed for several minutes at all negative potentials before the
channel ultimately would undergo an apparently spontaneous transition
to higher PO.
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Figure 6.
Spontaneous low-to-high
PO transition at negative potentials. A low
PO state at negative potentials was observed
transiently in approximately one-third of outward rectifiers in
inside-out patches. A, The two current traces on the
left represent an ORCC at 40 and 40 mV within the
first minute after channel activation (after a 30 sec, 60 mV voltage
pulse). Channel PO at 40 mV was 0.82. PO at 40 mV, in contrast, was 0.14. At low
PO, channel openings at 40 occurred
only in brief bursts, separated by long closures. The
right current traces represent the same channel at 40 and 40 mV 1 min later. PO at 40 mV was
0.87, which was only slightly higher than PO
at this potential 1 min earlier. PO at 40
mV, however, increased dramatically to 0.76. The long and short
closures of the channel at 40 mV now more closely resembled those at
40 mV above. The C adjacent to the dotted
line indicates the closed state of the channel.
B, Representative amplitude histograms of the
left and right current traces at 40 mV
shown in A. The small open peak (at 1.7 pA) in the
left histogram indicates a peak conductance of 44 pS.
The right histogram demonstrates both the increase in
PO at 40 mV and the increase in channel
conductance after this transition in PO.
Peak conductance was 51 pS. The black arrow pointing to
the right current trace in A calls
attention to the brief transition to the former peak conductance level
seen 1 min earlier. This conductance sublevel is represented by the 44 pS peak in the right histogram.
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We examined the open, closed, and burst time distributions of
representative ORCCs undergoing such a spontaneous transition from
low-to-high PO at negative potentials. This
analysis showed that spontaneous changes from low-to-high
PO (and experimental changes from high-to-low
PO shown later) arose from significant changes
in the long groups of channel openings, closures, and burst
distributions. The results from 26 spontaneous
PO transitions ("control" patches) are
summarized in Table 1. The experiments are described below.
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Table 1.
Open-state probability (PO) and
time constants from open time, closed time, and burst time
distributions
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Figure 6A shows current traces from a channel
undergoing a spontaneous transition from low-to-high
PO at negative potentials. Within the first
minutes of recording after activation, there was an obvious difference
in PO between the top left trace (40 mV) and the
bottom left trace (40 mV). In the top trace, channel PO was high (0.72). In the bottom trace, channel
PO was much lower (0.14). Within the next minute
of recording, however, the top right trace (40 mV) demonstrated
approximately the same high PO (0.77) as before,
but the bottom right current trace underwent a transition to higher
PO (0.68).
In Figure 6B, representative amplitude histograms of
current traces at 40 mV from Figure 6A are shown.
The small peak for open events at 1.7 pA (44 pS) demonstrates the low
PO of the channel at 40 mV. The increase in
PO at 40 mV 1 min later was accompanied by an
increase in channel conductance (from 44 to 51 pS). The phenomenon of
increased conductance with increased PO was
observed in many, but not all, patches that exhibited this early
transition from low-to-high PO. In addition,
very brief transitions to lower conductances also were seen sometimes
at positive potentials as well, although none was obvious in the
recording of the channel that is shown here. These results suggested
that the maximal conductance of the ORCC may be composed of
subconductances representing either molecularly distinct conducting
pores that function synchronously at high PO or
a single conducting pore with sublevels states of conductance rarely
evident at high PO.
Dwell time distributions
Visual inspection of channel activity in the right current traces
of Figure 6A, as well as channel activity in the
current traces presented earlier in Figure 1B,
suggested the presence of multiple closed states (Colquhoun and Hawkes,
1981). In Figure 7A, the right
histogram shows the distribution of closed intervals for the channel
shown in Figure 6A at 40 mV after high
PO is achieved. A second-order exponential fit
of this histogram indicated the presence of (at least) two closed
states. The two time constants for channel closed states,
S and L ("S" for short;
"L" for long) were 2.1 and 21 msec, respectively. For
26 patches, S was 3.0 ± 0.4, and L
was 21 ± 1.4. In the left histogram in Figure 7A, which demonstrates the closed time distributions of the low
PO state in the 40 mV current trace in Figure
6A, S was 2.5 msec, and
L was 85 msec, suggesting that the transition from
low-to-high PO arises in part from a decrease in
the time constant for long closures, whereas the time constant for
short closures remains primarily unchanged.
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Figure 7.
Closed, open, and burst time distributions at low
and high PO. A,
Left and right histograms represent the
distribution of closed events for the 40 mV traces of the channel
displayed in Figure 6. Both histograms were fit with second-order
exponentials. The two time constants displayed within the histograms
represent the mean times of the group of short closures
(S) and long closures (L).
For 26 control patches, mean S = 3.0 ± 0.4 msec
and L = 21 ± 1.4 msec. Thus, the spontaneous
transition from low-to-high PO in the
left and right current traces in
A involved primarily a change in the long group of
channel closures. B, Left and
right histograms graph the distribution of open events
at 40 mV before and after the transition in
PO. Second-order exponential fits suggested
two channel open states. For control patches, S = 3.6 ± 0.3 msec and L = 75 ± 3.9 msec
(n = 26). Thus, the low-to-high transition in
PO arose principally from a change in the
long open state. C, The left and
right histograms represent the distribution of channel
bursts at 40 mV for the transition between low and high
PO. As with open and closed states, the
burst distribution histograms could be fit accurately with a
second-order exponential. The shorter group of bursts
(1) and the longer group of bursts (2) were 121 ± 16 and 2777 ± 181 msec, respectively. Thus, the spontaneous transition from low-to-high
PO appears to arise mainly from an increase
in the longer group of bursts, although an increase in the short burst
lengths also was evident.
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The open time distributions of the channel are demonstrated analogously
in Figure 7B for the channel transitions from low-to-high PO at 40 mV shown in Figure 6. For the channel
at low PO, the short and long time
constants, S and L, from a
second-order exponential fit in the left histogram are 3.9 and 9.2 msec, respectively. The right histogram represents open time
distribution and a second-order exponential fit after the channel has
reached high PO. Now S is 4.2 msec, and L is 90 msec. For 26 patches at 40 mV,
S is 3.6 ± 0.3 and L is 75 ± 3.9. Thus, the increase in PO for the representative channel in Figure 6 appears to arise from an increase in
the time constant for long openings as well as a decrease in the time
constant for long closures. The time constant for short openings
remains primarily unchanged.
The short closed states also appeared to group channel openings into
bursts of activity. Figure 7C shows burst duration
histograms for the ORCC (from Fig. 6) as it underwent the transition
from low-to-high PO. Second-order exponential
fits indicated that burst durations could be described by two time
constants, 1 and 2, analogous to
the short and long time constants for closed and open states. At low
PO, 1 was 38 msec, and
2 was 254 msec for the channel in Figure 6. After 1 min,
at high PO, 1 was 71 msec and 2 was 2341 msec. For 26 patches showing high
PO activity, 1 and
2 were 121 ± 16 and 2777 ± 181 msec,
respectively, suggesting that the largest increase with the transition
to high PO was in the longer burst length.
Phalloidin and gelsolin modulate PO
and conductance
Phalloidin is a bicyclic heptapeptide "phallotoxin" derived
from poisonous mushrooms (Wieland and Faulstich, 1978) that binds actin
polymers much more tightly than actin monomers; it also shifts the
equilibrium between actin filaments and monomers toward filaments,
lowering the critical concentration for polymerization by 10- to
30-fold (Cooper et al., 1987). Stabilization of actin filaments arises
by reducing the subunit dissociation rate constants to near zero at
both ends of actin polymers (Pollard et al., 1990).
When excised inside-out patches were pulled in the presence of 10 µM phalloidin, the first obvious change was the
preservation of low PO at negative membrane
potentials. Figure 8A
illustrates activity from an ORCC activated in the presence of
phalloidin after the patch was stepped to 90 mV for 5 sec. At 40 mV
the mean PO was 0.70 ± 0.04 (n = 26) for channels in control solutions and
0.13 ± 0.03 (n = 11) in the presence of 5 µM phalloidin, representing a statistically significant
difference (p < 0.001). At 40 mV, however, the
mean PO was 0.81 ± 0.04 (n = 11) for control patches and 0.72 ± 0.04 (n = 11) in the presence of 5 µM phalloidin, representing a
statistically insignificant difference (p = 0.09).
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Figure 8.
Phalloidin and gelsolin modulate
PO and conductance. A,
Current traces from a patch excised in the presence of 5 µM phalloidin. In 11 patches exposed to phalloidin,
channels at negative potentials remained fixed at low
PO. B, A black
line is drawn from a short burst of channel openings to a
current trace that expands the short burst both in amplitude and time.
The top dotted line (labeled C)
represents the closed state of the channel. The next three dotted lines from top to bottom
represent three different channel subconductance levels. The current
trace on the right represents the same channel at 40
mV after the application (1 U/ml) of the actin-severing protein
gelsolin. Note the marked increase in channel PO. The peak amplitude of the current trace
on the right is indicated by the bottom dotted
line. This line also is reproduced as the lowest dotted
line in the left current trace before gelsolin
addition. C, Amplitude histograms plot the change in
conductance for the channel in the presence of phalloidin and after the
subsequent addition of gelsolin. Note the clear delineation of three
conductance levels (9, 24, and 35 pS) in the left
histogram. Peak conductance in the presence of phalloidin was 35 pS.
The right histogram shows the change in conductance
after the addition of gelsolin. Peak conductance increased to 44 pS
with increasing PO, and conductance sublevels were no longer evident.
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Table 1 compares mean channel open, closed, and burst times for the
ORCCs at 40 mV in the presence and absence of phalloidin. The
increase in long closed times (from 21 ± 1.4 to 97 ± 6.3 msec; p 
0.001), the decrease in long open times (from
75 ± 3.9 to 11 ± 1.1 msec; p 0.001), and
the decrease in long burst lengths (from 2777 ± 181 to 278 ± 39 msec; p 0.001) all contribute to the dramatic
decrease in PO at 40 mV in the presence of
phalloidin. Note that the time constants for the short open and closed
states and short burst lengths for phalloidin-treated patches were not significantly different from those in control solutions.
Preservation of the patch cytoskeleton with phalloidin not only lowered
PO but also allowed for the observation of
additional conductance sublevels otherwise not observed at high
PO. Figure 8B demonstrates
subconductance levels in the presence of phalloidin. The left histogram
in Figure 8C demonstrates the conductance sublevels as
individual peaks of current amplitude. Individual conductance sublevels
were observed in 11 patches excised in phalloidin, although the lowest
sublevel (9 pS) was difficult to resolve in seven of those patches.
Mean conductances for each sublevel were 8.8 ± 0.2, 25 ± 0.3, 35 ± 0.7, and 44 ± 0.6 pS (n = 11) in
patches treated with phalloidin.
Gelsolin is a Ca2+-dependent F-actin capping
and severing protein observed ubiquitously in all cells
(Vandekerckhove, 1990). In brain, it may be concentrated preferentially
in glial cells (Tanaka and Sobue, 1994). Gelsolin not only severs actin
filaments but also appears both to sever actin filaments in the
presence of phalloidin and possibly to displace phalloidin from actin
filaments. We examined the effects of 1 U/ml gelsolin applied to five
patches excised in the presence of phalloidin. As shown in the right
current trace in Figure 8B, application of gelsolin
resulted in a marked increase in channel PO and
in peak conductance level. Moreover, sublevels of conductance no longer
were observed.
A summary of the effects of phalloidin and phalloidin plus gelsolin on
channel conductance and gating is given in Figure
9. Cumulative data from 15 control, 7 phalloidin, and 5 phalloidin plus gelsolin experiments from which
complete I-V plots were obtained are illustrated in Figure
9. Note the dramatic decrease in PO at all
negative potentials for the phalloidin, as compared with the control
patches (membrane potentials >40 mV were excluded to avoid the
confounding influence of depolarization-dependent inactivation on
PO; see Fig. 2). Channel
PO after gelsolin treatment was not
significantly different from that under control conditions (p = 0.86).
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Figure 9.
PO and conductance
levels in the presence of phalloidin and phalloidin plus gelsolin.
These two graphs show the cumulative data from 15 control, 7 phalloidin, and 5 phalloidin plus gelsolin experiments from which
complete I-V plots were obtained. Membrane potentials
>40 mV were excluded to avoid the confounding influence of
depolarization-dependent inactivation on PO.
A, Plot of PO versus membrane
potential (mV) showing the decrease in PO at
negative potentials and the statistically insignificant decrease in
PO at positive potentials in the presence of
phalloidin (filled squares), as compared with
control patches (filled circles). The
actin-severing protein gelsolin (unfilled squares)
reverses the effects of phalloidin at negative potentials.
B, I-V plot reconstructed from the
single-channel data shown in Figure 8. In the presence of phalloidin,
mean conductance levels for all patches were 8.8 ± 0.2 (n = 4), 25.0 ± 0.3 (n = 9), 35 ± 0.7 (n = 8), and 44 ± 0.6 (n = 11). In this particular cell the peak level of
conductance at 40 mV was 35 pS (filled circles). After the addition of gelsolin, peak conductance
increased to 44 pS (unfilled circles). Each conductance
level within the negative voltage range was well fit by linear
regression via the expected reversal of the Cl
channel in symmetrical solutions. Dotted lines from the
regressions are extended into the positive voltage range to illustrate
the different degrees of I-V rectification at the level
of the single channel as it transitions between different conductance
sublevels at negative potentials.
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Figure 9B depicts current levels in the presence of
phalloidin and phalloidin plus gelsolin for the channel illustrated in Figure 8. Note that the I-V relationship for the sublevels
at negative conductances is well fit by linear regressions via the expected reversal of 0 mV. The significance of these regression fits
was extremely high: p < 0.005 for the 9 pS level,
p < 0.0001 for 24 pS level, and p < 0.005 for the 35 pS level. The fits were extended into the positive
voltage range to help in visualizing channel rectification with respect
to subconductance level. At negative potentials the conductance level
represented by the unfilled circles is well fit by linear regression
(p < 0.005).
Actin and cytochalasin D modulate PO at
negative potentials
The foregoing phalloidin-gelsolin experiments do not demonstrate
whether actin directly interacts with channels to change PO and conductance or whether actin works in
conjunction with other soluble modulators distinct from channel
proteins to affect channel function. We explored this issue by applying
actin directly to inside-out patches containing activated ORCCs. As
shown below, the addition of actin reduced PO
throughout the negative voltage range. This reduction was associated
with changes in the long open, closed, and burst time
distributions.
In Figure 10A, the
left current traces represent an outward rectifier after activation by
a 90 sec, 60 mV voltage pulse. At 40 mV (the top trace),
channel PO was 0.74; at 40 mV, channel PO was 0.71. These values represent
PO over 30 sec of recordings at each potential.
The right current traces demonstrate records of the same channel ~5
min after the addition of a mixed actin solution of polymers, short
filaments, and monomers (final concentration, 1 mg/ml). Actin caused a
profound decrease in PO at negative potentials, leaving PO at positive potentials unchanged. For
the channels in Figure 10, PO at 40 mV was
0.05 over 30 sec, whereas at 40 mV PO was 0.69 after the addition of actin. For seven patches exposed to actin
solutions, PO at 40 mV was 0.11 ± 0.04. This compares with the control PO of 0.70 ± 0.04 (n = 26). The changes in
PO for actin and control solutions are
summarized in Table 1. As with phalloidin, the changes in
PO appear to be the cumulative result of an
increase in the long closed state, a decrease in the long open state,
and a decrease in longer burst lengths.
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Figure 10.
Actin and cytochalasin D modulate
PO at negative potentials. A,
The left current traces are taken from a patch initially
in control solutions. PO at 40 mV was 0.74;
PO at 40 mV was 0.71. The
right current traces demonstrate records of the same
channel ~5 min after the addition of a mixed solution of actin
polymers, short filaments, and monomers (final concentration, 1 mg/ml). Actin caused a dramatic decrease in PO at
negative potentials. PO at 40 mV was 0.05 over 30 sec, whereas at 40 mV PO was 0.69 after the addition of actin. B, The current trace on the
left represents an outwardly rectifying channel at 40
mV after it had been exposed to 1 mg/ml actin.
PO at 40 mV was 0.23. The current trace on
the right shows the same channel 17 min after the
addition of 10 mM cytochalasin D while the amount of actin in the bath remained unchanged. Channel PO
increased to 0.95. Note also the marked increase in open channel noise
accompanying the increase in PO. The
C adjacent to the dotted line indicates the closed state of the channel. C,
PO is plotted versus membrane potential
(mV). Note that the increase in PO after the
addition of cytochalasin D (unfilled squares) surpasses
the PO of control patches. Mean
PO at 40 mV after the addition of
cytochalasin D was 0.89 ± 0.02 (n = 5),
significantly higher (p < 0.05) than the
PO (0.70 ± 0.04) in control
cells.
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In the presence of actin, application of cytochalasin D (Cyt D) to
excised patches caused a dramatic increase in PO
at all potentials (Fig. 10B). Excised patches,
however, required at least a 10-15 min incubation period before the
increase in PO was observed. Patches were
remarkably stable for prolonged periods after the addition of actin to
the recording bath. An example of an actin/Cyt D experiment is shown in
Figure 10B. The current trace at the left represents
an ORCC at 40 mV after it had been exposed to 1 mg/ml actin. The
PO of this channel at 40 mV in the presence of
actin is 0.23. The current trace on the right in Figure
10B shows the same channel 17 min after the addition
of 10 µM Cyt D while the amount of actin remains
unchanged. Channel PO in this case increased to
0.95 at 40 mV. Note also the marked increase in open channel noise
accompanying the increase in PO. Although this
noise potentially could arise from actin and/or cytochalasin directly
interacting with the channel pore, another explanation is that the open
channel noise is caused by ATP in the bath. ATP was always present as a
component of the actin solution. We previously observed a similar increase in open channel noise by directly adding ATP to activated channels (see Fig. 4B). Moreover, voltage-activated
channels in control experiments did not demonstrate any flickering or
open channel noise with Cyt D in the bath. The mean
PO at 40 mV after the addition of Cyt D was
0.89 ± 0.02 (n = 5), which was significantly higher (p < 0.05) than
PO (0.70 ± 0.04) in control patches at this potential. Control PO (0.81 ± 0.04)
and Cyt D PO (0.89 ± 0.02) at 40 mV were
not significantly different (p = 0.26).
A plot of PO versus membrane potential (mV) for
control, actin, and actin/Cyt D patches is shown in Figure
10C. Note the small but statistically significant
(p < 0.05) reduction in
PO observed at positive potentials. Mean
PO at 40 mV for control patches was 0.81 ± 0.02 (n = 26) whereas, for patches exposed to actin,
PO was 0.60 ± 0.03 (n = 7). Although actin may lower PO at positive potentials, the flickering block of ATP (see Fig. 4) also may explain
this reduction in PO, because ATP is a
component of the actin solutions. In Figure 10C, also note
the increase in PO at all negative potentials
after exposure to Cyt D. PO at all potentials in
the presence of cytochalasin exceeded even that measured in control
patches.
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DISCUSSION |
These results provide the first report of an ORCC in neocortical
astrocytes. Single-channel activation and kinetics suggest that this
Cl channel may contribute to morphology-dependent
and actin-dependent changes in astroglial whole-cell
Cl conductance (Lascola and Kraig, 1996).
Paralleling whole-cell currents, channel activation increased with
changes in cell morphology. In addition, single ORCCs were blocked
incompletely but irreversibly by DIDS, mimicking the DIDS inhibition of
whole-cell currents. Finally, depolarization-dependent inactivation was
observed in ORCCs, a kinetic feature of morphology-dependent whole-cell
Cl conductances. Voltage-dependent inactivation
was observed in single channels at a frequency (20-30%) close to that
observed in shape-dependent whole-cell currents (10-20%).
Results in this study also show that ORCCs can undergo marked changes
in PO and conductance spontaneously and that
these changes occur predominantly at membrane potentials negative to 0 mV. An analysis of open, closed, and burst time distributions between low and high PO states indicates that
spontaneous PO transitions arise from
significant changes in the longer groups of channel openings, closures,
and bursts. In the presence phalloidin, an agent that stabilizes
cytoskeletal actin, low PO states at negative potentials are preserved throughout the length of patch recordings. In
addition, channel conductance may be reduced with phalloidin, indicating that the outward rectifier has a greater tendency toward conductance sublevels at low PO. When phalloidin
patches were exposed further to the actin-severing protein gelsolin,
however, channel PO and conductance increased.
These results suggest strongly that spontaneous transitions in excised
patches from low-to-high PO and from lower to
higher conductance may arise from the gradual disassembly of
filamentous actin from the patch into the recording bath. Filamentous
actin appears to have a "restraining" or inhibiting effect on
astroglial ORCC activity, particularly within a range of negative
membrane potentials.
To support this hypothesis further, we found that the addition of actin
solutions to control patches lowered PO at
negative potentials. Again, changes in long openings, long closures,
and long bursts seem to account for the high-to-low transition in PO. These results provide strong evidence that
actin itself is required for the ORCC kinetic transitions and that
normally an intact cytoskeleton is not simply holding some soluble
inhibitor (e.g., a phosphatase) in place. Further evidence that actin
polymers, and not monomers, are responsible for the reduction in
PO follows from the subsequent application of
cytochalasin D to patches in the presence of actin. Cytochalasin D,
which shifts the equilibrium between actin polymers and monomers toward
monomers, leads to a dramatic reversal of the effects of actin. The
increase in PO at negative potentials actually
exceeds the PO of control cells. If actin
monomers in solution led to the initial decrease in
PO, cytochalasin D would not be expected
to influence channel PO.
Together, these results indicate that cytoskeletal actin can influence
the degree of rectification of whole-cell Cl
currents profoundly because of ORCC activation. Preservation of actin,
or the addition of actin to inside-out patches, leads to a 70-80%
decrease in channel PO at negative potentials,
whereas at positive potentials PO is reduced
only slightly, if at all. Moreover, at low
PO, the channel exhibits a tendency to
dwell at lower conductance sublevels. Combined, these phenomenon would lead to macroscopic currents with a much lower amplitude at negative potentials as compared with positive potentials (i.e., rectification would be greatly enhanced). Because astrocytes function only at negative membrane potentials in brain, cytoskeletal actin therefore would gate most of the physiological and pathological
whole-cell Cl conductance arising from ORCC
activation in these cells.
Actin and Cl channels in other cells
Several reports now exist from other cell preparations that
examine the interaction of single Cl channels with
actin and drugs that modulate actin. Two reports, one in muscle
(Haussler et al., 1994) and one in proximal tubule epithelia (Suzuki et
al., 1993), have investigated the interaction of actin with
Cl channels that resemble the astroglial ORCC. The
report in muscle cells is consistent with our observations, whereas the
study in proximal tubule cells provides contrary results. In the latter case, differences in experimental methods may explain the disparity of
their results with those in the muscle study and ours.
In the experiments performed in proximal tubule cells, Suzuki et al.
(1993) found that the application of cytochalasin D significantly decreased the mean open-state probability as well as the
number of active Cl channels in excised patches
from proximal tubule cells. If they added "long actin filaments"
back to patches after exposure to cytochalasin D,
Cl channel PO was restored
and the number of active channels increased. Cytochalasin D also
reduced whole-cell Cl currents in their
preparation. This result was in conflict with previous studies that
have shown that F-actin stabilization prevents Cl
secretion in epithelial cells (see Shapiro et al., 1991). Perhaps the
important difference between these studies and those of others as well
as ours is that Suzuki et al. (1993) examined PO
changes only within the first 6 sec after cytochalasin application. In both the muscle study and in our experiments, several minutes of
incubation with cytochalasins often were required before large changes
in PO were observed.
Haussler and colleagues (1994), in a study of the interaction between
actin and the muscle Cl channel, obtained results
more consistent with ours. Their principal observation was that
cytochalasin D activated whole-cell Cl currents
and increased the PO of previously activated
single Cl channels. An important similarity
between their work and ours is that PO for their
channel demonstrated a clear tendency to increase as membrane
potentials shifted positive. However, an important difference between
the two studies was that PO for their channel
appeared to remain fixed at nearly zero within the range of negative
voltages. In cell-free patches from astrocytes in our studies,
PO at negative potentials inevitably progressed
to values >0.50. Therefore, results of experiments in which
cytochalasins are applied directly to astroglial inside-out patches are
difficult to interpret, in that astroglial channel activity
spontaneously progressed to high PO without the
drug.
Perhaps astrocytes undergo a spontaneous transition to high
PO in control conditions, and muscle cells
remain fixed at a low PO because the cortical
actin cytoskeleton in these cells is markedly different. Muscle
cytoskeleton is extremely rigid. Filamentous actin in muscle has been
shown to be relatively resistant to depolymerization (Pollard et al.,
1990). This rigidity may arise in part because of the high
concentrations of actin-capping proteins in muscle (Vandekerckhove,
1990). Indeed, muscle myosin itself binds actin and adds stability to
actin polymers. Astrocytes, on the other hand, may have a more dynamic
cortical cytoskeleton. In fact, this would be consistent with the
observation that astrocytes possess a high capacity to undergo dramatic
morphological changes. Greater cytoskeletal flexibility would
facilitate this unusual capacity for morphological transformation.
Thus, the ease with which the astroglial cytoskeleton is poised to
undergo dynamic changes could manifest itself when membrane patches are
excised. With patch excision, actin may depolymerize spontaneously and diffuse into the recording bath. Spontaneous depolymerization of actin
would explain spontaneous low-to-high transitions in PO of the astroglial ORCC.
The actin cytoskeleton as a signal transducer in astrocytes
The results presented in this study suggest that cytoskeletal
actin may be part of a novel signaling mechanism in astrocytes that
regulates the activity of Cl channels. The
experiments in which actin was added directly to excised patches
provided compelling evidence that actin itself is interacting with
Cl channels or closely associated channel proteins
to modulate channel function. However, these experiments were not
intended to prove that only actin, and not other messenger systems,
could activate and modulate astroglial Cl
conductances. Instead, it is likely that actin is part of complex regulatory system in astrocytes and perhaps other cells that interacts with second messenger systems and target proteins to elicit changes in
membrane-based functions.
The functions of the almost 70 actin-binding proteins described to date
are regulated by second messengers, kinases, bioactive lipids, and
phospholipids (Stossel, 1989; Cooper, 1991; Matsudaira, 1991).
Ca2+, for example, regulates the cross-linking of
actin filaments by nonmuscle 
-actinins and activates the nucleating,
severing, and filament-capping activities of gelsolin, villin, and
severin. Analogously, phosphorylation by cAMP-dependent protein kinase A reversibly inhibits the bundling of actin filaments by synapsin I and
by erythrocyte band 4.9 (Stossel, 1989). Thus, changes in the physical
state of actin in astrocytes and other cells may arise from the complex
interplay of several messenger systems that act on the actin
cytoskeleton. If this is true, then cytoskeletal actin could be
considered a cellular "integrator." In astrocytes, actin appears to
integrate changes in cell form with changes in Cl
channel function.
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FOOTNOTES |
Received Oct. 8, 1997; revised Dec. 9, 1997; accepted Dec. 15, 1997.
R.P.K. was supported by a Grant from the National Institute of
Neurological Disorders and Stroke (NS-19108), a Zenith Award from the
Alzheimer's Association (ZEN-96-031), and the Brain Research Foundation of The University of Chicago. D.J.N. was supported by ROI
GM36823 and ROI GM54266 and a grant from the Cystic Fibrosis Foundation. C.D.L. was supported by an MD/PhD Training Grant in Growth
and Development (HD-07009) from the National Institute of Child Health
and Human Development and a National Research Service Award
(F31-MH11126) from the National Institute of Mental Health.
Correspondence should be addressed to Dr. Richard P. Kraig, Department
of Neurology, MC 2030, University of Chicago, 5841 South Maryland
Avenue, Chicago, IL 60637.
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Top
Abstract
Introduction
Compound via MeSH
