Cl
and
K+ conductances activated by cell
swelling in primary cultures of rabbit distal bright convoluted
tubules
I.
Rubera,
M.
Tauc,
C.
Poujeol,
M. T.
Bohn,
M.
Bidet,
G.
De
Renzis, and
P.
Poujeol
Unité Mixte de Recherche Centre National de la Recherche
Scientifique 6548, Université de Nice-Sophia Antipolis, O6108
Nice Cedex 2, France
 |
ABSTRACT |
Ionic currents induced by cell swelling were characterized in
primary cultures of rabbit distal bright convoluted tubule (DCTb) by
the whole cell patch-clamp technique.
Cl
currents were produced
spontaneously by whole cell recording with an isotonic pipette solution
or by exposure to a hypotonic stress. Initial
Cl
currents exhibited
outwardly rectifying current-voltage relationship, whereas steady-state
currents showed strong decay with depolarizing pulses. The ion
selectivity sequence was I
= Br
> Cl
glutamate. Currents
were inhibited by 0.1 mM 5-nitro-2-(3-phenylpropylamino)benzoic acid
and 1 mM 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid and strongly blocked by 1 mM diphenylamine-2-carboxylate. Currents were
insensitive to intracellular Ca2+
but required the presence of extracellular
Ca2+. They were not activated in
cells pretreated with 200 nM staurosporine, 50 µM
LaCl3, 10 µM nifedipine, 100 µM verapamil, 5 µM tamoxifen, and 50 µM dideoxyforskolin.
Staurosporine, tamoxifen, verapamil, or the absence of external
Ca2+ was without effect on the
fully developed Cl
currents. Osmotic shock also activated
K+ currents in
Cl
-free conditions. These
currents were time independent, activated at depolarized potentials,
and inhibited by 5 mM BaCl2. The
activation of Cl
and
K+ currents by an osmotic shock
may be implicated in regulatory volume decrease in DCTb cells.
whole cell; cell volume; ion conductance; kidney
 |
INTRODUCTION |
IN MANY EPITHELIAL CELLS,
Cl
channels are essential
for the transport of salt and water across the membrane bilayer, for
stabilization of the resting membrane potential, and for regulation of
cell volume. At least three distinct
Cl
currents, regulated by
adenosine 3',5'-cyclic monophosphate (cAMP), cytosolic
Ca2+, and osmotic pressure, have
been found in several tissues, including airway epithelial cells, sweat
gland, pancreas, and T84 intestinal cells (14). In the kidney,
Cl
channels activated by
cell swelling have been observed in proximal tubule, in the loop of
Henle, and in collecting tubules (39). As in other tissues, these
channels are one of the membrane transport pathways that mediate an
efflux of osmolytes to bring about regulatory volume decrease (RVD).
The activation of Cl
condutance during RVD is regularly associated with an increase in
K+ conductance, leading to a net
loss of KCl and a concomitant efflux of water. In the proximal tubule,
cell swelling is mainly the result of the transport of osmotically
active solutes into the cells via cotransporters coupled to the
Na+ gradient. In these cells,
mechanisms of RVD are well developed and have been extensively studied
(19, 22, 23, 25, 41, 44). Presently, very few studies have examined
swelling-activated conductances in the distal convoluted tubule,
although, with the low osmolarity of the fluid delivered to this
segment, the distal cells would be expected to require volume
regulatory capabilities. In view of these indications, we carried out
whole cell experiments to investigate further the hypotonically
activated Cl
and
K+ conductances in distal
convoluted tubule. For this purpose, we used primary cultures of the
bright part of rabbit distal convoluted tubule (DCTb). These cultures
are now well characterized (26). They exhibit apical
Cl
channels activated by
cAMP and basolateral
Ca2+-dependent
Cl
conductance (3, 35, 43).
This study describes hypotonically activated
Cl
currents with
pharmacological properties similar to those of swelling-activated
Cl
current in epithelial
cells undergoing RVD on exposure to hyposmotic medium. The activation
of this Cl
conductance
requires extracellular Ca2+ and is
associated with the phosphorylation of P-glycoprotein. The hypotonic
shock also activates K+
conductance sensitive to Ba2+. It
is therefore possible that the observed
K+ and
Cl
currents may be
responsible for RVD.
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MATERIAL AND METHODS |
Primary Cultures
The primary cell culture technique used in this study has been
described in detail in previous papers (26, 35). Briefly, the bright
parts of the rabbit distal tubules were microdissected under sterile
conditions from kidneys obtained from 4- to 5-wk-old male New Zealand
rabbits. Kidneys were perfused with Hanks' solution (GIBCO) containing
600-700 kU/l collagenase (Worthington) and cut into small
pyramids, which were incubated in medium containing 150 kU/l
collagenase. The tubules were seeded in collagen-coated 35-mm petri
dishes filled with a culture medium composed of equal quantities of
Dulbecco's modified Eagle's medium and Ham's F-12 (GIBCO),
containing 15 mM NaHCO3, 20 mM
N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), pH 7.5, 2 mM glutamine, 5 mg/l insulin, 50 nM
dexamethasone, 10 µg/l epidermal growth factor, 5 mg/l transferrin,
30 nM sodium selenite, and 10 nM triiodothyronine. Cultures were
maintained at 37°C in a 5%
CO2-95% air water-saturated
atmosphere. The medium was changed 4 days after seeding and then every
2 days.
Whole Cell Experiments
Whole cell currents were recorded from 12- to 22-day-old cultured cells
grown on collagen-coated supports maintained at 33°C throughout the
experiments. The ruptured-patch whole cell configuration of the
patch-clamp technique was used. Patch pipettes were made from
borosilicate capillary tubes (1.5 mm OD, 1.1 mm ID; Clay Adams) using a
two-stage vertical puller (PP 83; Narishige, Tokyo, Japan). When filled
with N-methyl-D-glucamine chloride (NMDG-Cl) solution the pipettes had a resistance ranging from 2 to 3 M
in a
NMDG-Cl buffer. An Ag-AgCl pellet was used as the reference electrode.
To reduce junction potentials, this electrode was bathed in an
identical solution to that contained in the pipette and connected to
the bath via a 3 M KCl-agar bridge. Cells were observed by using an
inverted microscope (Zeiss IM 35), the stage of which was equipped with
a water robot micromanipulator (MHW30, Narishige). The patch pipette
was connected via an Ag-AgCl wire to the head stage of a RK 400 patch
amplifier (Biologic). After the formation of a gigaohm seal, the fast
compensation system of the amplifier was used to compensate for the
head-stage intrinsic input capacitance (
8 pF). The membrane was
ruptured by additional suction to achieve the conventional whole cell
configuration. At this stage, the cell capacitance
(Cm) was
compensated for with a facility provided on the RK 400 amplifier. With
NMDG-Cl in the bath and in the pipette, the
Cm of 146 cells
was 27.2 ± 0.8 pF (mean ± SE), and the series resistance
(Rs) averaged
9.8 ± 0.8 M
. With potassium glutamate in the bath and
in the pipette, the
Rs averaged 6.5 ± 0.3 M
(n = 32).
Rs was
compensated for in K+ currents but
not in Cl
currents. For this purpose, the
Rs compensation
circuitry of the RK 400 was used. This facility allowed a 90%
compensation, reducing the capacitive time constant in the same
proportion. However, in all cases, experiments where
Rs > 20 M
were discarded. Cell membrane potentials were measured at zero membrane
current in the current-clamp mode of the amplifier. Extracellular test solutions were perfused into the bath using a four-channel glass pipette, the tip of which was placed as near as possible to the clamped
cell.
Voltage-clamp commands, data acquisition, and data analysis were
controlled by an IBM-AT-compatible computer equipped with a Digi Data
1200 interface (Axon Instruments, Foster City, CA). Commercially
available pCLAMP software (version 6.0, Axon Instruments) was used to
generate whole cell current-voltage
(I-V) relationships. Membrane
currents resulting from voltage stimuli were filtered at 1 kHz, sampled
at a rate of 2,560/s, and stored directly onto the hard disk. For the
measurement of chloride currents, cells were held at a holding
potential
(Vhold) of
50 mV, and 400-ms pulses from
100 to +120 mV were applied
with increments of 20 mV every 2 s.
K+ currents were measured by
applying pulses ranging from
100 to +80 mV from a
Vhold of
50 mV.
Fluorescence Experiments
Image analysis. The optical system was
composed of a Zeiss ICM-405 inverted microscope and a Zeiss ×40
objective, which was used for epifluorescent measurements with a 75-W
xenon lamp. The excitation beam was filtered through narrow-band
filters (340, 360, and 380 nm; Oriel) mounted in a motorized wheel
(Lambda 10-2; Sutter Instrument) equipped with a shutter to
control the exposure times. The incident and the emitted fluorescence
radiation were separated through a Zeiss chromatic beam splitter.
Fluorescence emission was selected through a 510-nm narrow-band filter
(Oriel). The transmitted light images were viewed by an intensified
camera (Extended ISIS; Photonic Science, Sussex, UK). The 8-bit
extended-ISIS camera was equipped with an integration module to
maximize signal-to-noise ratio. The video signal from the the camera
proceeded to an image processor integrated in a DT2867 image card (Data
Translation) installed in a Pentium 100 PC. The processor converts the
video signal into 512 lines by 768 square pixels per line by 8 bits per
pixel. The 8-bit information for each pixel represents one of the 256 possible gray levels, ranging from 0 (for black) to 255 (for white).
Image acquisition and analysis were performed by the 2.0 version of AIW
software (Axon Instruments). The final calculations were made using the
Excel software (Microsoft).
Intracellular
Ca2+
measurements. Fifteen- to twenty-day-old confluent DCTb
monolayers grown on petri dish were loaded with a solution of 2 µM
fura 2 containing 0.01% pluronic acid for 30 min at 37°C and were
then washed with a NaCl solution. The cells were successively excited
at 340 and 380 nm; the images were digitized and stored on the hard
disk of the computer. Each raw image was the result of an integration
of six frames averaged four times. The acquisition rate was 1 image/4
s. The intracellular Ca2+
concentration was calculated from the dual-wavelength fluorescence ratio by using the Grynkiewicz equation (16).
Mn2+
influx measurements. Fifteen- to twenty-day-old
confluent DCTb monolayers were loaded for 45 min with a solution of 5 µM of the acetoxymethyl ester of fura 2 (fura 2-AM) containing 0.01% pluronic acid, and the cells were then washed with NaCl solution. Fluorescence quenching experiments were carried out by addition of 50 µM of MnCl2 in the NaCl medium.
Fluorescent measurements were performed at 360 nm (isobestic point of
fura 2) where the fluorescence signal is independent of free calcium
concentration. The manganese influx was quantified by the slope of the
quenching kinetics. Each raw image was the result of an integration of
six frames averaged four times. The acquisition rate was 1 image/4 s.
Cell volume estimation. The relative
cell volume was monitored by measuring trapped dye concentrations. This
technique was derived from that previously reported (44), but fura 2 was used instead of
2',7'-bis(carboxyethyl)-5(6)-carboxyfluorescein. The cells
were loaded with fura 2, as described above, and the fluorescence was
monitored with 360-nm excitation wavelength. At 360 nm, the variations
of the signal emitted by the probe are directly proportional to the
variations of the cell volume. In a typical experiment, the cells were
first perfused with an isotonic NaCl solution containing (in mM) 110 NaCl, 5 KCl, 1 CaCl2, 90 mannitol,
and 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid [pH 7.4, osmotic pressure
(Posm) = 320 mosmol/kgH2O at 30 ml/min],
and images were averaged 30 times and recorded every 5 s for 4 min.
Once the fluorescence stabilized, a hypotonic shock was induced by
perfusing the NaCl solution without mannitol
(Posm = 227 mosmol/kgH2O). The estimation of
relative change in cell volume from the fluorescent signal was made,
assuming that a 30% decrease in the osmolarity caused a decrease of
the fluorescent signal corresponding to a maximum swelling of 30%
compared with the initial volume. The means of relative volume changes
were obtained by the analysis of 5-10 zones in each of
n number of cultures chosen with the
software. Each zone delimited a cytoplasmic area chosen in individual
cells.
Drugs
A stock solution of 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB)
from Calbiochem was prepared at 100 mM in dimethyl sulfoxide (DMSO) and
used at 0.1 mM in final solutions. Diphenylamine-2-carboxylate (DPC)
from Aldrich was prepared as 1 mol/l stock solution in DMSO and
dissolved at 1 mM in incubation medium.
4,4'-Diisothiocyanostilbene-2,2'-disulfonic acid (DIDS)
from Sigma was directly dissolved at a final concentration of 1 mM.
Fura 2-AM from Molecular Probes was dissolved at 3 mM in DMSO.
Staurosporine and protein kinase inhibitor were from Calbiochem.
Nifedipine, verapamil, dideoxyforskolin, and tamoxifen were obtained
from Sigma.
Solutions
The compositions of the different solutions used in these experiments
are given in Table 1.
 |
RESULTS |
Cl
Currents in Unstimulated Cultured
DCTb Cells
Whole cells currents were recorded with
Ca2+-free pipette solutions
containing NMDG-Cl (Table 1, solution f ) and in
an extracellular solution containing NMDG-Cl (Table 1,
solution c). Both solutions were
isotonic (290 mosmol/kgH2O). After
successful gigaohm seal formation, the whole cell configuration was
obtained in 25% of the cases. Within 2-4 min after the mechanical
rupture of the membrane, the cell depolarized as the pipette solution
equilibrated with the cell interior. Voltage-clamp experiments were
performed, holding the cell at
50 mV and applying voltage steps
of 400-ms duration every 2 s from
100 to +120 mV in 20-mV
increments. Once cell depolarization was complete, currents elicited by
the voltage protocol were recorded every minute. Figure
1,
A-D, shows families of currents
recorded immediately, 2, 4, or 6 min after the beginning of whole cell
recording (recording time). Only 105 of 265 cells displayed these
currents in isotonic conditions. In these cells, we have considered
that the control currents (t = 0 min)
were those recorded when membrane potential just reached 0 mV. In these conditions, the initial currents measured 6 ms after the onset of the voltage pulse, rectified slightly in the
outward direction (Fig. 1E). They
reversed at
2.3 ± 1.4 mV
(n = 20), and the total current at 100 mV was 1.9 times the current at
100 mV (100 mV = 257.4 ± 24.4 pA,
100 mV =
137.1 ± 14.9 pA;
n = 20, P < 0.01). The initial currents then
increased with time and stabilized after 6 min (Fig. 1,
B-D).The
I-V relationships for initial and
steady-state currents are illustrated in Fig. 1,
E and
F, respectively. Initial currents
displayed an outward rectification that increased with time (Fig.
1E). The maximal current was reached
6 min after the beginning of the whole cell recording. At this time,
the initial current recorded at 100 mV was 2.4 times the current at
100 mV (100 mV = 1,117 ± 77 pA,
100 mV =
474.1 ± 73.0 pA; n = 20, P < 0.001). These large, outwardly
rectifying currents showed time-dependent inactivation at depolarizing
step potentials >40 mV. Generally, the time course of this
inactivation could be well fitted with a single exponential regardless
of the recording time. In 15 of the 20 cells studied, the greater the
voltage, the faster was the rate of the decay. At 120 mV, the currents
inactivated with a time constant of 132.4 ± 6.50 ms
(n = 15). To better illustrate the
decay of the currents, the percentage of inactivation was calculated as
the difference between the initial current at 6 ms and the steady
current at 390 ms into the same voltage step divided by the current at
390 ms. At every imposed potential, the inactivation was independent of
the recording time. Figure 2 shows that the
inactivation during the 400-ms pulse >40 mV strongly increased with
more depolarizing potentials. These data were obtained from experiments
performed in symmetrical Cl
concentrations. The reversal potential
(Erev)
was very close to that of
Cl
, and, in the absence of
permeable cations in the pipette, the outward current was carried by
Cl
.

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Fig. 1.
Spontaneous Cl currents in
unstimulated cultured rabbit distal bright convoluted tubule (DCTb)
cells. Membrane voltage was held at 50 mV and stepped to test
potential values between 100 mV and +120 mV in 20-mV increments
(A,
inset). Pipette and bath contained
N-methyl-D-glucamine chloride (NMDG-Cl)
solutions (solutions f and
c in Table 1). Whole cell currents
were recorded at t = 0 (A),
t = 2 (B),
t = 4 (C), and
t = 6 min
(D) after the mechanical rupture of
membrane. E and
F: average current-voltage
relationships measured 6 and 390 ms, respectively, after onset of
pulse. Values are means ± SE of 20 cells from 5 monolayers: ,
control (t = 0); , 1 min; , 2 min; , 3 min; , 4 min; , 6 min.
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Fig. 2.
Effect of depolarizing potentials on decay of
Cl currents. Pipette and
bath contained NMDG-Cl solutions
(solutions c and
f, Table 1). Percentage of
inactivation was calculated from ratio of initial current at 6 ms to
steady current at 390 ms into same voltage step. Each value represents
mean ± SE of 74 cells.
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To study the anion permeability of the cell membrane, all except 2 mM
of the Cl
in the bath
solution was replaced with
I
,
Br
, or glutamate. Figure
3,
A-C, gives typical recordings of
the currents obtained at 6 min in the presence of the three different anions. Figure 3, D and
E, shows
I-V relations for initial and steady-state currents. Table 2
summarizes Erev
values as well as the calculated permeability ratios obtained for a
given anion. To minimize the effects of capacitance transients, the
whole cell currents were fitted to an exponential curve over the
interval of 20-390 ms. The amplitude of the instantaneous current
was extrapolated to the time of the onset of the voltage step. In the
presence of I
or
Br
,
Erev shifted
toward the negative values and was independent from the duration of the
stimulation. Replacing external chloride with glutamate markedly
reduced the initial and steady-state outward currents. During this
substitution, the inward currents carried by chloride remained
unaffected, and
Erev moved toward
positive voltages. However,
Erev values
calculated for the initial I-V curves
were significantly lower than those calculated for steady-state currents. The curve in Fig. 4 illustrates
the variations of
Erev for
glutamate as a function of the voltage pulse duration. Finally, the
sequence for this conductance was
I
= Br
> Cl
glutamate.

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Fig. 3.
Effect of extracellular Cl
substitution on spontaneous
Cl currents in unstimulated
cells. Pipette contained NMDG-Cl solution
f. A-C: whole
cell currents recorded in the presence of
I
(solution a),
Br
(solution b), and glutamate
(solution d), respectively.
D and
E: average current-voltage
relationships measured 6 and 390 ms after the onset of the pulse,
respectively. Each value represents mean ± SE; , control in 140 mM NMDG-Cl (15 cells from 8 monolayers); , 140 mM NaI (7 cells from
3 monolayers); , 140 mM NaBr (6 cells from 3 monolayers); , 140 mM Na glutamate (7 cells from 2 monolayers).
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Fig. 4.
Extrapolated reversal potential of currents recorded in the presence of
external glutamate plotted as a function of duration of voltage pulse.
Pipette contained NMDG-Cl (solution
f), and the bath contained 140 mM Na glutamate
(solution d). Values are means ± SE of 7 cells from 2 monolayers.
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To further characterize the
Cl
current, we tested three
anion channel blockers added separately to the bathing solution. Figure 5,
A-D, gives typical traces of the
current obtained 6 min after starting the whole cell recording. The
addition of NPPB, DIDS, or DPC to the bathing solution inhibited the
whole cell Cl
currents
within 2 min. The effects of these blockers were reversible on washing
(data not given). The I-V
relationships for initial and steady-state currents are given in Fig.
5, E and
F, respectively. Overall, 0.1 mM NPPB
or 1 mM DIDS inhibited reversibly both initial inward (%inhibition at
100 mV: NPPB, 44.3 ± 3.8, n = 8; DIDS, 34.6 ± 5.0, n = 18) and
outward currents (%inhibition at +100 mV: NPPB, 71.4 ± 4.6, n = 8; DIDS, 77.2 ± 6.8, n = 18). The effects of NPPB and DIDS
were therefore voltage dependent. By contrast, the blocking effects of
DPC on initial currents were similar at
100 and +100 mV
(%inhibition at
100 mV, 71.3 ± 2.8; %inhibition at +100
mV, 73.5 ± 4.6, n = 5).

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Fig. 5.
Effect of Cl channel
inhibitors on spontaneous
Cl currents in unstimulated
cells. Pipette and bath contained NMDG-Cl solutions
(solutions c and
f ). A: whole
cell currents recorded at maximum of the response.
B: with extracellular perfusion of
10 4 M
5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB).
C: with extracellular perfusion of
10 3 M
4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS).
D: with extracellular perfusion of
10 3 M
diphenylamine-2-carboxylate (DPC). E
and F: average current-voltage
relationships measured 6 and 390 ms, respectively, after onset of
pulse. Values are means ± SE: , control (31 cells from 9 monolayers); , NPPB (8 cells from 3 monolayers); , DIDS (18 cells
from 4 monolayers); , DPC (5 cells from 2 monolayers).
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Cl
Currents Induced by a Hypotonic
Shock
Currents recorded in these DCTb cells were produced spontaneously when
the monolayers were bathed with a solution having the same osmotic
pressure as the pipette solution. The overall characteristics of this
Cl
conductance are similar
to the swelling-induced currents described in several tissues (8, 9,
21, 40, 49, 52). To study the effects of changes in osmotic pressure on
the development of this conductance, currents were then induced by
osmotic shock. In these experiments, the pipette solution was
maintained at 290 mosmol/kgH2O.
Moreover, to eliminate any participation of cations in the inward
current, experiments were carried out after replacing Na+ in the bath solution by
NMDG+. Figure
6A
illustrates the time course of the initial currents measured at +100 mV
as a function of the osmolarity of the bath solution. At an
extracellular solution osmolarity of 350 mosmol/kgH2O (Table 1,
solution c + 60 mM mannitol), the
voltage-step protocol elicited small, time-independent currents that
changed linearly with the membrane voltage, with a slope conductance of
0.97 ± 0.04 nS and
Erev of
0.69 ± 0.03 mV (n = 21).
Because of their small amplitude, the nature of these currents was not
analyzed further. The monolayer was then perfused with a 290 mosmol/kgH2O solution. In >95%
of the cells, an increase in the whole cell current was observed within
1 min. The currents reached a maximal after 5-6 min and remained
stable for 5 min. When the cells were reexposed to hyperosmotic
solution, the currents returned to the control level within 2-3
min. The currents induced by hypotonicity were analyzed at their
maximal values. It was found that they were very similar to those
developed spontaneously in isotonic solutions. This is illustrated by
the I-V relationships for initial currents reported in Fig. 6B. The
substitution of external Cl
by I
increased the outward
currents and moved
Erev toward more
negative values, giving a
Erev of
21.6 ± 0.94 mV and a calculated relative I
-to-Cl
permeability ratio of 2.40 ± 0.11 (n = 5). Furthermore, glutamate caused
Erev to shift by
38.5 ± 1.2 mV such that the glutamate-to-chloride permeability
ratio was 0.20 ± 0.01 (n = 5).
Figure 6B also shows the effect of
DIDS and NPPB on swelling-activated chloride currents. The blocking
effect of both drugs was voltage dependent. However, the potency of
NPPB to decrease the inward current was found to be greater than that
of DIDS (%inhibition at
100 mV: NPPB, 68.0 ± 7.1; DIDS,
17.9 ± 3.6; %inhibition at +100 mV: NPPB, 80.1 ± 8.2, n = 5; DIDS, 78.1 ± 6.4, n = 5).

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Fig. 6.
Cl currents induced by
hypotonic shock. A: time course of
initial current at 100 mV measured 6 ms after onset of pulse as a
function of bath osmolarity. B:
average current-voltage relationships measured 6 ms after the onset of
pulse. , Control current at 290 mosmol/kgH2O (21 cells from 5 monolayers); , current in hyperosmotic (350 mosmol/kgH2O) solution (21 cells
from 5 monolayers); , 140 mM NaI (5 cells from 2 monolayers); ,
140 mM Na glutamate (5 cells from 2 monolayers); , DIDS,
10 3 M (5 cells from 2 monolayers); , NPPB, 10 4
M (5 cells from 3 monolayers). Values are means ± SE.
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Regulation of the Cl
Conductance
Induced by Hypotonic Shock
Calcium. To eliminate the implication
of cytosolic Ca2+ in the
development of hypotonicity-induced
Cl
currents, experiments
were generally performed using pipette solutions containing 5 mM
ethylene glycol-bis(
-aminoethyl
ether)-N,N,N',N'-tetraacetic acid (EGTA) without additional
Ca2+. However, some experiments
were carried out by increasing EGTA concentration up to 10 mM. In 100%
of the cells tested, this maneuver did not modify the
Cl
currents on exposure to
a hypotonic medium (data not given). In further experiments, the
putative variations of cytosolic
Ca2+ induced by hypotonic shock
were followed by using fura 2 as a Ca2+ probe. The data clearly
indicated that no significant variation of intracellular
Ca2+ could be detected even with a
hypotonic shock of 100 mosmol/kgH2O (isotonicity:
intracellular Ca2+ concentration,
83.7 ± 6.0 nM; hypotonicity: intracellular
Ca2+ concentration, 79.3 ± 6.3 nM; n = 13).
The effects of extracellular Ca2+
on the development of hypotonicity-induced
Cl
currents were also
tested. The histogram of Fig. 7 shows that when the hypotonic shock was carried out in the absence of bath Ca2+, the development of the
Cl
current was
significantly impaired. In fact, in the absence of extracellular
Ca2+, 7 of the 13 recorded cells
did not exhibit Cl
currents
in response to a hypotonic shock, whereas the remaining six cells
elicited a response, which was reduced by 46.2 ± 8.1% compared with the control value. Conversely, removal of bath
Ca2+ after the current had been
activated did not modify significantly the amplitude of the
Cl
currents in any of the
cells tested (Fig. 8). These experiments indicate that extracellular Ca2+
was required to activate the swelling-activated
Cl
conductance. To
determine whether this activation was related to
Ca2+ influx across the cell
membrane, we studied the role of ionomycin in the development of the
Cl
conductance. To provide
evidence that ionomycin induced an increase of divalent cation entry,
we took advantage of the property of fura 2 fluorescence being quenched
by Mn2+, a commonly used
substitute for Ca2+. The
fluorescence at 360 nm (isobestic point for fura 2) is independent of
cytosolic Ca2+. Thus only fura 2 quenching due to Mn2+ uptake by
the cells modified the fluorescence signal. When NaCl solution
containing MnCl2 was perfused, an
immediate fluorescence decrease (0.30 ± 0.01 fluorescence arbitrary
units/s, n = 3) was observed. As shown
in Fig. 9, extracellular application of
ionomycin accelerated the rate of quenching (0.73 ± 0.03 fluorescence arbitrary units/s, n = 3), indicating an enhanced influx of divalent cations. This uptake
mechanism was blocked by 10 µM lanthanum (data not given). In a
second series of experiments, the effects of ionomycin were tested on
whole cell Cl
currents
developed in the presence or the absence of intracellular free
Ca2+. To avoid the apparition of
swelling-induced Cl
conductance, the currents were recorded in monolayers continuously perfused with hypertonic NMDG solution (Table 1,
solution c + 60 mM mannitol). In the
experiments shown in Fig.
10A,
whole cell currents were recorded in the absence of EGTA in the pipette
solution (Table 1, solution f without
EGTA). After control macroscopic currents were recorded, 2 µM
ionomycin was added to the bathing NMDG solution, and the stimulated
currents were recorded after 1 min. In these conditions, the
cytoplasmic free Ca2+ rose at 1.00 ± 0.19 µM (n = 13). Figure
10A shows that in the presence of
ionomycin, the currents increased during depolarizing voltage pulses.
The kinetics of the macroscopic current were clearly time dependent for
depolarizing potentials with a slow-developing component. The
corresponding I-V relationships for
early and steady-state activated currents are given in Fig.
10A. Currents reversed at
0.25 ± 0.2 mV (n = 6). Instantaneous
currents showed a slight outward rectification: the inward current at
100 mV was 263.7 ± 41.0 pA, and the outward current at +100
mV was 353.7 ± 34.0 pA. The steady-state current presented marked
outward rectification with an inward current at
100 mV of 214.0 ± 74.0 pA and an outward current at +100 mV of 524.0 ± 78.0 pA
(n = 6). In the experiments of Fig.
10B, whole cell currents were recorded
in the presence of 5 mM EGTA in the pipette solution (Table 1,
solution f). In 12 of the 20 cells
analyzed, the addition of 2 µM ionomycin induced the activation of
Cl
currents within 3-4
min. Under these conditions, ionomycin also enhanced the
divalent-cation influx across the membrane (see above), but this
increase was not accompanied by visible variations of intracellular
Ca2+ (data not given). The
iononycin-activated Cl
currents showed time-dependent inactivation at depolarizing step potentials of >60 mV and displayed an outwardly rectified
instantaneous I-V plot (Fig.
10B) with an
Erev of
0.35 ± 0.01 mV (n = 12).
Overall, these currents were quite similar to those induced by
hypotonic shock.

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Fig. 7.
Regulation of Cl
conductance induced by hypotonic shock. Treatment of cells with
the various compounds was carried out prior to exposure to osmotic
shock. Initial currents were recorded at 100 mV, 6 ms after the onset
of pulse. Control currents were recorded in hyperosmotic solution
(hyper) and after the hypotonic shock (21 cells from 7 monolayers).
Currents were then triggered in cells pretreated with 200 nM
staurosporine (staur, 4 cells from 2 monolayers), 10 µM protein
kinase inhibitor (PKI, 8 cells from 3 monolayers), 50 µM
La3+ (8 cells from 4 monolayers);
10 µM nifedipine (nife, 5 cells from 2 monolayers), 100 µM
verapamil (vera, 7 cells from 3 monolayers), 5 µM tamoxifen (tamo, 8 cells from 4 monolayers), 50 µM dideoxyforskolin (ddfk, 3 cells from
2 monolayers). Cl currents
were also recorded in the absence of external
Ca2+ (13 cells from 5 monolayers).
Values are means ± SE.
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Fig. 8.
Regulation of Cl
conductance induced by hypotonic shock. Treatment of cells with the
various compounds was carried out after osmotic shock, once
Cl currents were fully
developed. Control currents were recorded in hyperosmotic solution and
after the hypotonic shock (15 cells on 13 monolayers). Currents were
then triggered in cells pretreated with 200 nM staurosporine (staur, 3 cells from 3 monolayers), 100 µM verapamil (vera, 3 cells from 3 monolayers), 5 µM tamoxifen (tamo, 5 cells from 3 monolayers), 50 µM dideoxyforskolin (ddfk, 3 cells from 2 monolayers).
Cl currents were also
recorded in the absence of external
Ca2+ (4 cells from 4 monolayers).
Values are means ± SE.
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Fig. 9.
Fura 2 fluorescence quenching by external manganese: effect of
ionomycin. Entry of manganese into cytoplasm was monitored by
fluorescence quenching measurements at 360 nm in fura 2-loaded cultured
DCTb. Initial decrease was the result of perfusion of NaCl buffer
containing 50 µM MnCl2. After a
control period of 2 min, 2 µM ionomycin was added to the bath.
Decrease of fluorescence induced by ionomycin was compared with that of
control period. Values are means ± SE of 10 cells from 1 monolayer. Identical experiments were repeated in 3 different
monolayers.
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Fig. 10.
Cl currents induced by
extracellular ionomycin in hypertonic NMDG solution
(solution c + 60 mM mannitol).
A: whole cell currents were recorded
in the absence of EGTA in the pipette solution
(solution f without EGTA). Average
current-voltage relationships were measured 6 ( ) and 390 ms ( )
after onset of pulse. Values are means ± SE of 6 cells from 3 different monolayers. B: whole cell
currents were recorded in the presence of 5 mM EGTA in pipette solution
(solution f). Average
current-voltage relationships were measured 6 ( ) and 390 ms ( )
after onset of pulse. Values are means ± SE of 12 cells from 6 different monolayers.
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The development of hypotonicity-induced
Cl
currents was further
studied in the presence of several factors that could interfere with
Cl
conductance.
Lanthanum. In a series of experiments,
hypotonic shock was carried out in the presence of 50 µM
La3+ in the bath solution. This
trivalent cation irreversibily suppressed the
Cl
currents (Fig. 7).
P-glycoprotein-related inhibitors. To
investigate a possible involvement of P-glycoprotein in the generation
of the hypotonicity-induced Cl
currents, inhibitors for
this protein were tested. As shown in Fig. 7, verapamil (100 µM) and
nifedipine (10 µM) completely prevented activation of
Cl
currents on exposure of
the cells to a hypotonic solution. Similar inhibition was also induced
by the extracellular application of tamoxifen (5 µM). These
inhibitors have been known to reverse multidrug resistance by
inhibiting P-glycoprotein-mediated drug efflux and to block the
P-glycoprotein-associated
Cl
channel (53). The
effects of these drugs were partially reversed while rinsing the
monolayers with a solution without inhibitors. Verapamil and tamoxifen
were also applied separately following preactivation of
Cl
currents by hypotonic
shock. As illustrated in Fig. 8, neither drug significantly affected
the fully developed Cl
currents. Dideoxyforskolin has also been reported to inhibit swelling
Cl
conductance thought to
be related to the expression of P-glycoprotein. As shown in Fig. 7, 50 µM dideoxyforskolin prevented activation of the current on hypotonic
swelling. Moreover, the drug also produced a marked irreversible
inhibition (68.4 ± 4.8%, n = 3) of the preactivated swelling-induced Cl
current (Fig.
8).
Intracellular ATP. All the experiments
were performed with 5 mM MgATP in the pipette solution. However, to
check the influence of cytosolic ATP on the generation of
swelling-induced Cl
currents, a series of experiments was carried out in the absence of ATP
in the pipette medium. Under these conditions, exposure of the cells to
hypotonic solutions induced small
Cl
currents, the amplitude
of which slowly decreased and stabilized after 6-8 min (initial
currents recorded at 100 mV 6-8 min after hypotonic shock in the
presence of ATP, 1,937.0 ± 328.9 pA,
n = 5; in the absence of ATP, 541.7 ± 34.7 pA, n = 5).
Protein kinases. To check
whether activation of the
Cl
current by hypotonic
stress could be regulated by protein kinases, the effects of inhibitors
of protein kinase A and protein kinase C were studied. When the
hypotonic shock was carried out in the presence of 10 µM of protein
kinase A inhibitor (PKI) in the pipette solution, the whole cell
Cl
currents developed
normally in all cells tested (Fig. 7). When the shock was performed in
the presence of 200 nM staurosporine in the bath medium, no significant
increase in Cl
currents was
recorded (Fig. 7). However, bath staurosporine did not affect the
magnitude of the Cl
currents once they were preactivated by the hypotonic solution (Fig.
8).
K+ Currents
Induced by Hypotonic Shock
The effect of hypotonic swelling was tested under
Cl
-free conditions. The
first experimental series was carried out with 140 mM potassium
glutamate and EGTA in the pipette solution and 140 mM sodium glutamate
in the bath medium. In each experimental condition, only positive
current curves were observed. Figure
11A
illustrates the family of current recordings made in a hypertonic bath
medium with test potential that ranged from
100 mV to +80
mV in increments of 20 mV. Outward currents were
significantly different from 0 at
60 mV (23.9 ± 11.0 pA;
n = 25, P < 0.05) and increased during more
positive voltage pulses. These currents showed virtually no
inactivation during the 400-ms pulse. The corresponding
I-V curve in Fig.
11E confirms that the channels
involved in this conductance were activated at depolarized potentials.
The conductance measured by the maximal slope of the
I-V curve averaged 24.0 ± 2.2 nS
(n = 25). As illustrated in Fig.
11B, outward currents were activated when the hypertonic bath solution was replaced by an isotonic solution
of identical ionic composition. Maximal current activation was reached
2 min after the onset of the osmotic shock. The
I-V curve of Fig.
11E shows that the activation of
outward currents became significantly different from control from
40 mV (control current, 53.6 ± 16 pA; volume-activated
current, 111.0 ± 10 pA; P < 0.01, n = 25) and increased with
depolarizing potentials. The calculated maximal slope conductance
increased significantly (34.5 ± 2.7 nS;
P < 0.01, n = 25) compared with control values (see above). Whole cell conductance was then analyzed following the
addition of 5 mM Ba2+ to the
isotonic bath solution. Figure 11, C
and E, clearly indicate that, at
Vhold between
40 mV and +20 mV, the currents recorded in the presence of
Ba2+ were significantly lower than
those recorded in isotonic bath solution without
Ba2+. Conversely, the currents
determined for depolarizing voltage steps were not significantly
altered. Experiments in the control bathing solution following this
intervention showed that the effect of
Ba2+ could be completely reversed
(data not given). Finally, reexposing the same cells to hypertonic
solution induced a decrease in the amplitude of the outward currents
within 1 min (Fig. 11C). The current
amplitude returned to values slightly lower than that initially
measured (Fig. 11D), suggesting that
currents generated by osmotic shock were associated with cell swelling.
At negative imposed potentials, block of the outward conductance by
Ba2+ strongly indicated that the
currents induced by the osmotic shock were mainly the result of an
efflux of K+ from the cell.
Moreover, in the absence of permeable anions in the solutions, it is
very likely that the outward current measured at positive potentials
was also carried by K+. To further
analyze the ionic nature of these conductances, a series of experiments
was performed whereby all Na+ in
the bathing medium was replaced by
K+. Under these conditions,
osmotic shock increased both inward and outward currents (Fig.
12A)
with a Erev close
to 0 mV. The initial current recorded at 80 mV was measured to be 3.6 times the current at
80 mV (80 mV = 2,600 ± 232 pA;
80 mV =
718 ± 94 pA;
n = 9, P < 0.001). These large, outwardly
rectifying currents were time independent and had a maximal slope
conductance of 40.2 ± 2.6 nS (n = 9). They were carried by K+, which
was the only permeable ion into pipette and bath solutions. To confirm
the implication of a K+
conductance, the effect of Ba2+ in
the bathing medium was then tested. The recording of Fig. 12B and the
I-V plot of Fig.
12E show that 5 mM
BaCl2 almost completely blocked
the inward current without significantly modifying the outward current.
After the washout of Ba2+, the
inward conductance recovered completely (Fig.
12C). Finally, reexposure of cells
to a hypertonic solution inhibited both outward and inward currents
(Fig. 12, D and
E).

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Fig. 11.
K+ currents induced by a hypotonic
shock in the presence of external
Na+. Membrane voltage was held at
50 mV and stepped to test potential values between 100 mV
and +80 mV in 20-mV increments. Pipette contained 140 mM K glutamate
(solution g). Whole cell currents
were successively recorded in hyperosmotic Na glutamate solution
(A), in isosmotic Na glutamate
solution (B), in the presence of 5 mM BaCl2
(C), and in hyperosmotic Na
glutamate solution (D).
E: average current-voltage
relationships: , control in hyperosmotic Na glutamate solution; ,
isosmotic Na glutamate solution; ,
BaCl2, 5 mM; , hyperosmotic Na
glutamate solution. Values are means ± SE of 30 cells from 21 monolayers.
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Fig. 12.
K+ currents induced by a hypotonic
shock in the presence of external
K+. Membrane voltage was held at
50 mV and stepped to test potential values between 100 mV
and +80 mV in 20-mV increments. Pipette contained 140 mM K glutamate
(solution g). Whole cell currents
were successively recorded in isosmotic K glutamate solution
(A), in the presence of 5 mM
BaCl2
(B), after rinsing out
BaCl2
(C), and in hyperosmotic K glutamate
solution (D).
E: average current-voltage
relationships: , isosmotic K glutamate solution; ,
BaCl2, 5 mM; , hyperosmotic K
glutamate solution. Values are means ± SE of 9 cells from 3 monolayers.
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Influence of Hypotonic Shock on Relative Cell Volume
Figure
13A
shows that the reduction of the osmolarity of the perfused solution
caused a rapid increase in relative cell volume. This cell swelling was
followed by a RVD. One minute after the hypotonic shock, the relative
cell volume reached 128.9 ± 0.8% (n = 9 cells from 3 monolayers) of the
initial volume and returned to 106.1 ± 0.3% of the original volume
within 2 min. To test the involvement of
K+ conductance in the RVD process,
experiments were carried out in the presence of 5 mM external
Ba2+. When
Ba2+ was perfused with the
hypotonic solution, the cells never returned to their initial volume
(Fig. 13B). To demonstrate that
Cl
conductance was involved
in RVD of distal cells, the effect of the
Cl
channel blocker NPPB was
then tested. Figure 13C shows that,
when 0.1 mM NPPB was added to the hyposmotic solution, RVD was
completely abolished.

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Fig. 13.
Effect of a hypotonic shock on cultured DCTb. After loading with fura
2, cultures were rinsed in a 300 mosmol/kgH2O NaCl solution, and
fluorescence of fura 2 was measured at 360 nm for a control period of 2 min. Then a hypotonic shock was induced by perfusing a 200 mosmol/kgH2O NaCl solution. Images
were recorded every 5 s. After analysis, relative volume change in
percent of initial volume was plotted against time.
A: normal hypotonic solution
(n = 9).
B: hypotonic solution with 5 mM
BaCl2
(n = 6).
C: hypotonic solution with 0.1 mM NPPB
(n = 16). Identical experiments were
repeated in 3 different monolayers.
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DISCUSSION |
In the present study, we have demonstrated that cultured DCTb cells
exhibit swelling-activated whole cell
Cl
currents. These currents
are clearly distinct from the
Ca2+-activated
Cl
currents and
cAMP-dependent Cl
currents
we have previously described in these primary cultures (3, 35, 43). In
the first series of experiments presented here, spontaneously
developing Cl
currents were
measured when isotonic bathing and pipette solution were used. The
currents reached a maximum 6-7 min after establishment of the
whole cell configuration. According to Worrell et al. (51), this
increase of Cl
conductance
could be the result of cell swelling in isosmotic medium. Under these
conditions, the presence of a diffusion barrier within the cytoplasm
could impair the complete equilibration between the cell interior and
the pipette solution. Consequently, the intracellular medium would
maintain a significant oncotic pressure due to the conservation of
high-molecular-weight proteins. In a second series of experiments,
Cl
currents also developed
in cells exposed to hypotonic shock. Control currents were obtained by
maintaining the cells in a bath, which was 50 mosmol/kgH2O hypertonic to the
pipette solution. In 95% of cells tested, the currents were of very
small amplitude and time independent. The measured
Erev was close to
0 mV, indicating that most of the control current was carried by
Cl
ions. The remaining 5%
of cells developed swelling
Cl
currents immediately
after the whole cell configuration was obtained. This was probably
because the hypertonicity of the bath medium was not sufficient to
prevent cell swelling. In the hypertonic solution, the basal
Cl
currents remained small
and almost constant for at least 8 min. When the osmolarity of the bath
solution was lowered to 290 mosmol/kgH2O, whole cell
conductance increased ~10-fold within 6-8 min. Once maximally
developed, the current remained very stable and could be rapidly
inhibited by reexposing the cells to the hypertonic bath solution.
The biophysical and the pharmacological characteristics of the
Cl
conductances induced
either spontaneously in isotonic medium or induced by hypotonic stress
show strong similarities with the properties of swelling-activated
Cl
currents described in
many other epithelial cells (8, 9, 21, 40, 49, 52). The peak currents
exhibited an outwardly rectifying I-V
relationship, whereas the steady-state currents showed a strong decay
at depolarizing pulses. This decay was time dependent and increased
with increasing cell depolarization. As previously suggested (47), this
phenomenon may well correspond to the closure of channels already
activated at the holding potential. Nevertheless, the physiological
relevance of this inactivation is questionable, since the current
inactivation was obtained at voltages far removed from the
Cl
equilibrium potential.
The swelling-induced currents were carried mainly by
Cl
. This was confirmed by
the removal of extracellular
Cl
, which strongly reduced
the amplitude of the outward currents. Moreover, when the major cations
in the pipette and bath solutions were replaced with NMDG,
Erev remained
close to zero, as in symmetrical chloride solutions. This finding
clearly demonstrates that the major part of the swelling-induced
current was Cl
selective.
However, when glutamate was substituted for
Cl
in the bathing solution,
the Erev was
shifted to just 60 mV. Because the theoretical
Erev for
glutamate substitution was >100 mV, the
Cl
channel could be
slightly permeable to glutamate. Low levels of organic anion
permeability have already been reported in kidney cells (48).
Interestingly,
Erev for
glutamate was dependent on the duration of the stimulation. Thus the
relative permeability for glutamate was higher when calculated using
instantaneous currents than using steady-state conductance. Two
hypotheses could be proposed to support this observation:
1)
Cl
and glutamate could
permeate the cell membrane through separate channels both being
activated by hypotonicity. The channels activate at depolarizing
potential, but the conductance deactivation kinetic could be faster for
glutamate than for Cl
currents. Volume-sensitive anion channels with a relative high permeability to glutamate, aspartate, and taurine were already found in
Madin-Darby canine kidney (MDCK) cells (2). However, according to the
recent findings of Boese et al. (4), the hypotonic stress would
activate a common pathway for conductive
Cl
and amino acids (as
glutamate or taurine). 2)
Cl
and glutamate fluxes
process through the same channel. It is possible that large
depolarizing potentials modify the relative permeability at the onset
of the voltage pulse. For example, depolarization could induce
important changes in channel conformation (54), resulting in transitory
modifications in the channel selectivity.
The halide selectivity sequence permits the different types of
Cl
channels to be
identified. In DCTb cells, the sequence for the hypotonicity-induced
Cl
current was
I
= Br
> Cl
. In many cell types
studied, swelling-induced
Cl
channels have been
reported to be more permeable to
I
than to
Br
(6, 49). However, this
is not a constant finding, because, in some tissues, the anion
conductivity sequence was found identical to that we reported in the
present study (8). Sensitivity to various anion channel blockers also
helps to distinguish the type of
Cl
channel under
investigation. For this purpose, we examined the effects of NPPB, DPC,
and DIDS. NPPB and DIDS modified mainly the outward current, whereas
DPC strongly blocked both outward and inward currents. In the
literature, the voltage dependence of the blocking effect of DIDS and
the voltage independence of the inhibitory effect of both DPC and NPPB
are common characteristics of swelling-activated
Cl
currents. In cultured
DCTb cells, NPPB acted in a similar fashion to DIDS, suggesting that
their blocking mechanisms may be identical. However, it is necessary to
note than the effect of DIDS on the inward current was much more
constant from one cell to another than the effect of NPPB.
Despite abundant literature, the precise mechanism underlying the
activation of swelling-induced
Cl
current remains unclear.
In several studies, an increase in cytosolic Ca2+ has been reported during the
hypotonic stress (24, 25). In the present study, we found that the
activation of a swelling-induced Cl
conductance took place
in the presence of high concentrations of EGTA in the pipette solution.
Moreover, we also demonstrated that no significant cytosolic
Ca2+ variation could be detected
during the hypotonic shock. These results demonstrate that DCTb
Cl
conductance during
swelling is probably insensitive to intracellular Ca2+. This finding agrees with
that reported for a number of cells, including human sweat gland
(11), endothelial (31), epididymal (8), intestinal 407 (21), and T84 cells (51). Moreover, in proximal convoluted tubules
(PCT), the changes in cytosolic Ca2+ were shown to play no role in
RVD and, therefore, on swelling-induced activation of ion conductances
(5). Nevertheless, other studies have underlined a positive role of
cytoplasmic Ca2+ in the control of
the volume-sensitive Cl
currents (12, 24). The question of whether
Ca2+ is involved in the activation
of these currents is not so clear cut. Interestingly, we have
demonstrated that the removal of external Ca2+ just before the hypotonic
shock completely impaired
Cl
current activation,
suggesting that Ca2+ influx could
participate in activating the
Cl
channels. The
observation that the Ca2+ channel
blocker La3+ also abolished the
Cl
current activation
corroborates this hypothesis. However, to better support this
hypothesis, we studied the effects of ionomycin in the activation of
Cl
channels in hypertonic
bathing medium. The ionophore induced two types of
Cl
conductances, depending
on the presence of high EGTA concentrations in the pipette solution. In
the absence of chelator, the
Cl
currents induced by
ionomycin were roughly identical to the
Ca2+-dependent
Cl
currents previously
found in cultured DCTb (3). These currents were likely stimulated by an
increase in cytoplasmic Ca2+ due
to Ca2+ release from intracellular
stores. By contrast, in the presence of EGTA, the application of
ionomycin increased the uptake of Mn2+. This increase is evidence of
Ca2+ uptake across the membrane
(7). This maneuver induced large Cl
currents, which
inactivated during depolarizing voltage steps. Although they are not
fully characterized, these currents closely resemble the
swelling-activated Cl
currents described in the present study. Finally,
Ca2+ entry across the plasma
membrane could be one of the mediators of the activation of
Cl
currents by
hypotonicity. The role of external
Ca2+ was already postulated by
McCarty and O'Neil (25). They showed that RVD in proximal straight
tubules was highly dependent on the extracellular
Ca2+ concentration. Moreover,
they concluded that Ca2+ channels
may be responsible for a swelling-activated
Ca2+ entry. In previous papers, we
have shown that Ca2+ channels are
present in the apical membrane of DCTb cells (34) and may represent the
apical influx pathway for transepithelilal Ca2+ transport. These
Ca2+ channels are distinct from
the nonselective cation channels and are blocked by nifedipine,
verapamil, and La3+. In the
present study, the problem has been to better characterize the
involvement of the calcium channels in the control of swelling-induced conductance. Nifedipine and verapamil are the only drugs that block the
Ca2+ channel in DCTb (36), and
these drugs also directly interfere with the swelling
Cl
channels (see below).
While the response to hypotonic shock was developing, removal of
external Ca2+ did not affect the
Cl
current. Very similar
results were previously found in pancreatic duct cells (49), whereon
the authors concluded that Ca2+ is
only involved in events occurring early in the mechanism. Their
conclusions also support our data.
In most of the experiments described here, ATP was included in the
pipette solutions. When ATP was omitted from the pipette, hypotonic
stress always triggered Cl
conductance, but the currents never reached their maximum amplitude and
rapidly decreased with time. It is therefore probable that this
decrease began as the endogenous ATP was washed out after the whole
cell configuration was established. This characteristic of the
swelling-induced Cl
current
is similar to that reported previously in several tissues (11, 31, 49).
However, divergent results were also obtained in other cells (18, 40).
One of the hypotheses advanced to explain the action of ATP is that
swelling-induced Cl
currents are associated with the multidrug-resistance P-glycoprotein. Thus the gradual depletion of endogenous ATP would decrease the activity of this protein.
To determine whether P-glycoprotein is implicated in the
swelling-induced Cl
conductance of DCTb cells, we have tested the effect of different drugs
that have been reported to inhibit
Cl
currents thought to be
related to the expression of P-glycoprotein (46, 47).
Verapamil, nifedipine, tamoxifen, and dideoxyforskolin strongly
impaired the development of swelling currents. However, tamoxifen and
verapamil failed to produce significant inhibition of preactivated
Cl
current, whereas
dideoxyforskolin almost completely supressed it. Taken together, these
findings indicate that P-glycoprotein could participate in the
regulation of the swelling-induced
Cl
current but that this
protein is probably not itself the actual Cl
channel. This
observation supports recent literature reports of different cell types
(17, 45).
P-glycoprotein is a drug transporter modulated by protein kinase C
(PKC). We therefore examined whether PKC might also play a role in the
control of the swelling-activated
Cl
current. Bath
application of staurosporine, which is a membrane-permeable inhibitor
of PKC, completely inhibited the development of the swelling-induced
conductance. However, staurosporine never suppressed the current when
applied after the commencement of the hypotonic chock. These results
agree with those obtained for pancreatic duct cells (49) and suggest
that PKC regulates swelling-induced Cl
currents in cultured
DCTb. If it is generally accepted that PKC is implicated in the control
of this conductance, the exact nature of this control is under
discussion. In fact, according to some authors, activation of PKC
inhibits the channel by phosphorylation of P-glycoprotein (17), whereas
others have shown that PKC increases the channel activity (49) or is
not involved in the mechanism of regulation (42, 45). On the basis of
data obtained in pancreatic duct cells, Verdon et al. (49) have
proposed a very attractive hypothesis. They suggest that cell swelling
leads to an influx of Ca2+ and
that the concomittant increase of intracellular
Ca2+ will be sufficient to
activate PKC. The PKC in turn phophorylates a regulatory protein. This
protein could be the P-glycoprotein. Such an intracellular signaling
pathway was proposed by Nishizuka (32) in a recent review, at least up
to the point activation of the PKC.
It remains to be shown how intracellular
Ca2+ can increase in the presence
of a high EGTA concentration. The observations of Evans and Marty (10)
shed light on this problem by indicating that, with EGTA as a buffer, a
whole region of the cell could escape control by the
Ca2+ buffer. Because this region
could extend to a large part of the plasma membrane (10), a local
transient increase of Ca2+ could
arise without being detected by fluorescence methods.
Because activation of the PKA pathway is known to regulate
Cl
channels in epithelia
(14), we therefore investigated this possibility on the
swelling-induced Cl
conductance of DCTb cells. The use of the specific PKA inhibitor peptide, PKI (5, 24), clearly indicates that PKA is not
involved in activating the channel. This finding is strengthened by
experiments in which we demonstrated that the application of forskolin
did not modify the activation of the
Cl
conductance during the
hypotonic shock (data not given). In epithelial cells, the activation
of the swelling-induced Cl
conductance generally appears to be independent of cAMP (31, 49).
In many cells, swelling induced by exposure to hyposmotic solutions is
followed by RVD mediated by KCl loss via
K+ and
Cl
channels. Concerning the
mammalian kidney, RVD has been primarily studied in PCT (19, 22, 23,
25, 41, 44), in medullary thick ascending limb cells (30), and in the
cell line MDCK (1, 20). In a previous study (22), we demonstrated that
primary cultures of PCT responded to hypotonic shock by activating
K+ and
Cl
conductance. Further
experiments on rabbit proximal straight tubules brought evidence for a
swelling-sensitive Cl
conductance in the basolateral membrane (23, 37). Swelling-induced Cl
channels were also
described in the distal nephron, notably in RCCT-28A cells, an
immortalized cell line derived from rabbit collecting duct (39) and
recently in primary culture of rat inner medullary collecting duct
cells (50) and in M-1 cell line obtained from mouse collecting duct
(29). Interestingly, the Cl
channel recorded in RCCT-28A cells is regulated by a signaling pathway
that involves PKC and could mediate
Cl
efflux during RVD. To
determine whether cultured DCTb cells develop RVD after a hypotonic
shock, we used a simple fluorescence method for studying relative cell
volume variations (44). The findings indicate that DCTb cells are
sensitive to osmolarity changes in the bathing medium and that they are
capable of RVD after a hypotonic shock. The RVD process was impaired by
NPPB, confirming the implication of a
Cl
-conductive pathway.
Moreover, the observation that
Ba2+ completely blocked RVD
strongly indicates the involvement of K+ conductance. The first
experiments that we have performed to characterize cation conductance
during osmotic challenge indicate that
K+ currents also increased during
the hypotonic shock. To characterize this
K+ conductance, it was necessary
to use Cl
-free solutions,
because, in the presence of KCl, the swelling-activated current was
mainly due to Cl
(data not
given). In the presence of symmetrical
K+ solutions, the whole cell
conductance activated by the osmotic shock was
Ba2+ sensitive.
Ba2+ blocked mainly the inward
currents, indicating a strong voltage dependence as is expected for
Ba2+.
Ba2+-sensitive
Ca2+-dependent maxi
K+ channels have been found in the
apical membrane of cultured proximal tubules (28), cultured cortical
ascending limb of the loop of Henle (27), and cortical collecting duct
cells (13, 15). However, in the present experiments, the pipette
solution contained 5 mM EGTA, and it is unlikely that
Ca2+-activated maxi
K+ currents participated in the
resting K+ conductance. As
discussed above for Cl
channels, the osmotic shock could modulate
Ca2+ influx through
Ca2+ channels, and a local
increase of cytosolic Ca2+ could
activate the Ca2+-sensitive maxi
K+ channel. A similar mechanism
has already been postulated to explain the RVD in primary cultures of
renal proximal tubule (19, 22, 44).
It is of interest to examine the effect of osmotic stress on
Cl
and
K+ conductances determined close
to the resting membrane potential. With consideration to an imposed
potential of
60 mV 2 min after the beginning of the osmotic
shock, the outward Cl
current was increased by 56% (initial current at
t = 0 min,
88.2 ± 9.0 pA;
initial current at t = 2 min,
157.8 ± 17.0 pA; n = 19, P < 0.01),
whereas the outward K+ current was
not significantly modified. An increase in outward K+ current became significant at
40 mV (see RESULTS).
Therefore, cell swelling induced an increase predominantly in
Cl
current. Such increase
has also been demonstrated in rat colonic crypt (9) and rat mesangial
cells (33). According to these studies, hypotonic cell swelling could
induce a depolarization of membrane potential. If it is also the case
in our experiments, the activation of a swelling-dependent
Cl
conductance might
depolarize the membrane potential toward a value where the efflux of
K+ from the cell becomes
significant. Finally, in cultured DCTb, a hyposmotic shock could well
induce a net loss of KCl, thus allowing the cells to regulate their
volume.
In summary, DCTb cells in primary culture exhibit hypotonic
shock-induced Cl
and
K+ conductances and RVD process.
The Cl
currents described
here share characteristics with the P-glycoprotein Cl
currents described
in other systems (31, 46, 52), although P-glycoprotein is more likely
to be a regulator of the channel than the chloride channel itself
(17). The accompanying K+ currents
are strongly voltage dependent and blocked by external Ba2+. Thus the activation of
Cl
and
K+ currents by an osmotic shock
may be implicated in regulatory volume decrease in DCTb cells.
 |
FOOTNOTES |
Address for reprint requests: P. Poujeol, UMR-CNRS 6548, Bâtiment Sciences Naturelles, Université de Nice-Sophia
Antipolis, Parc Valrose, 06108 Nice Cedex 2, France.
Received 20 December 1996; accepted in final form 18 June 1997.
 |
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