Unité Mixte de Recherche Centre National de la Recherche
Scientifique 6548, Université de Nice-Sophia Antipolis, 06108 Nice Cedex 2, France
The role of cystic fibrosis transmembrane
conductance regulator (CFTR) in the control of Cl
currents was studied in mouse kidney. Whole cell clamp was used to
analyze Cl currents in primary cultures of proximal and
distal convoluted and cortical collecting tubules from wild-type (WT)
and cftr knockout (KO) mice. In WT mice, forskolin activated
a linear Cl current only in distal convoluted and
cortical collecting tubule cells. This current was not recorded in KO
mice. In both mice, Ca2+-dependent Cl
currents were recorded in all segments. In WT mice, volume-sensitive Cl currents were implicated in regulatory volume decrease
during hypotonicity. In KO mice, regulatory volume decrease and
swelling-activated Cl current were impaired but were
restored by adenosine perfusion. Extracellular ATP also restored
swelling-activated Cl currents. The effect of ATP or
adenosine was blocked by 8-cyclopentyl-1,3-diproxylxanthine. The
ecto-ATPase inhibitor ARL-67156 inhibited the effect of hypotonicity and ATP. Finally, in KO mice, volume-sensitive Cl
currents are potentially functional, but the absence of CFTR precludes
their activation by extracellular nucleosides. This observation
strengthens the hypothesis that CFTR is a modulator of ATP release in epithelia.
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INTRODUCTION |
THE CYSTIC FIBROSIS
TRANSMEMBRANE CONDUCTANCE REGULATOR (CFTR) protein has been
detected by electrophysiological techniques in a variety of cultured
cells of the renal tubule, such as distal convoluted tubule (DCT)
(21), cortical collecting tubule (CCT) (2,
30), and inner medullary collecting duct (10). In
these segments, the presence of CFTR is correlated with activation of a
cAMP-activated Cl current. However, along the nephron,
CFTR is not always associated with these Cl currents. For
instance, despite the presence of CFTR transcripts, CFTR expression,
along with forskolin-induced conductance, was not detected in rabbit
proximal tubule in primary culture (21). This observation
highlights the fact that CFTR could play an important role in the
control of different channels in kidney tissue. Such control is now
well established in secretory epithelia. In these structures, besides
the cAMP-sensitive Cl secretion, CFTR controls the
epithelial Na+ channel (12, 14, 18, 28) and
the outwardly rectifying Cl channel (24).
Moreover, CFTR is also needed for an effective volume regulation in
airway and intestinal epithelia (31, 32), suggesting that
it could modulate K+ and Cl channels
implicated in regulatory volume decrease (RVD). Indeed, these multiple
functions of CFTR could explain the different phenotypes induced by
cystic fibrosis (CF) in secretory epithelia. In contrast, the role of
CFTR in the kidney remains uncertain, inasmuch as there is no major
disruption of renal function in CF patients (27).
Nevertheless, the reduced renal excretion of NaCl observed in CF
indicates that Cl and Na+ channels could be
dependent on CFTR expression, suggesting that mutation of CFTR could
induce a primary defect in renal function. A better understanding of
the function of CFTR in the kidney, therefore, seems to be necessary.
For this reason, we chose to investigate the role of CFTR along the
nephron using primary cultures of proximal convoluted tubules (PCT),
DCT, and cortical collecting ducts microdissected from the kidney of
cftr
/ and cftr+/+ mice. The
cftr/ mice lack cAMP-activated
Cl currents in the colon, airways, and exocrine pancreas
cells (6) and represent a useful model for studying the
different ion channel defects due to CF. In the present study, using
patch-clamp methodology, we confirmed that cAMP-sensitive
Cl conductances measured in primary cultures of DCT and
cortical collecting tubule (CCT) cells are linked to CFTR integrity.
Moreover, in contrast to the data reported in the literature on airways and endothelial cells (33), an increase in
Ca2+-dependent Cl channels does not
compensate for the lack of CFTR Cl channels in renal
tissue. The PCT, DCT, and CCT cells from
cftr/ mice lost their capacity to regulate
their volume after a hypotonic shock because of the impairment of
swelling-activated Cl channels. In
cftr/ cells, the activity of these channels
could be restored by external application of adenosine. This suggests that CFTR controls the swelling-activated Cl channels by
modulating adenosine autocrine production in renal cells.
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MATERIALS AND METHODS |
Animals
Knockout CFTR mice were generated with the gene-targeting
methodology previously described (26) at Centre de
Développement des Techniques Avancées pour
l'Expérimentation Animale (Orléans, France). This
strain of mice was originally derived from ES129/Sv cells injected into
C57BL/6 embryos. They were backcrossed with C57BL/6 mice for three
generations and then intercrossed. Mice were allowed free access to
food and water in a facility at 25 ± 1°C with a 12:12-h
light-dark cycle. The 4- to 6-wk-old wild-type cftr+/+ mice
and cftr/ mice homozygous for the disrupted
cftr gene were killed by cervical dislocation, and the
kidneys were removed. All experiments were performed in accordance with
the guidelines of the French Agricultural Office and the legislation governing animal studies.
Primary Cell Cultures
PCT, DCT, and collecting tubules were microdissected under
sterile conditions. Kidneys were perfused with Hanks' solution (GIBCO)
containing 700 kU/l collagenase (Worthington), cut into small pyramids
that were incubated for 1 h at room temperature in perfusion
buffer (160 kU/l collagenase, 1% Nuserum, and 1 mM CaCl2),
and continuously aerated. The pyramids were then rinsed thoroughly in
the same buffer devoid of collagenase. The individual nephrons were
dissected by hand in this buffer under binoculars using stainless steel
needles mounted on Pasteur pipettes. The criteria used to identify the
nephron segments have been described elsewhere (4).
Briefly, PCT corresponded to the 1- to 1.5-mm segment of tissue located
immediately following the glomerulus. The DCT portion was the segment
between the macula densa and the first branching with another tubule
[i.e., connecting tubule (CNT)]. The CNT segment was discarded. The
CCT was identified as the straight, poorly branched portion that
followed the CNT segment. After they were rinsed in dissecting medium,
tubules were transferred to collagen-coated 35-mm petri dishes filled
with culture medium composed of equal quantities of DMEM and Ham's
F-12 (GIBCO) containing 15 mM NaHCO3, 20 mM HEPES, pH 7.4, 1% serum, 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 removed 4 days after seeding and then every 2 days.
Electrophysiological Studies
Whole cell currents were recorded from 6- to 20-day-old cultured
cells grown on collagen-coated supports maintained at 33°C for the
duration of the experiments. The ruptured-patch whole cell
configuration of the patch-clamp technique was used. Patch pipettes (2- to 3-M
resistance) were made from borosilicate capillary tubes (1.5 mm OD, 1.1 mm ID; Popper Manufacturing) using a two-stage vertical
puller (model PP 83, Narishige, Tokyo, Japan) and filled with a
solution containing (in mM) 140 N-methyl-D-glucamine (NMDG) chloride, 1 or 5 EGTA, 5 MgATP, and 10 HEPES, pH 7.4. The bath solution contained (in
mM) 140 NMDG chloride, 1 CaCl2, 60 mannitol, and 10 HEPES, pH 7.4. Cells were observed using an inverted microscope; the
stage of the microscope was equipped with a water robot
micromanipulator (model WR 89, Narishige). The patch pipette was
connected via an Ag-AgCl wire to the head stage of a patch amplifier
(model RK 400, Biologic). After formation of a gigaseal, the
fast-compensation system of the amplifier was used to compensate for
the intrinsic input capacitance of the head stage and the pipette
capacitance. The membrane was ruptured by additional suction to achieve
the conventional whole cell configuration. Settings available on the amplifier (model RK 400) were used to compensate for cell capacitance. No series resistance compensation was applied, but experiments in which
the series resistance was >20 M were discarded. Solutions were
perfused in the extracellular bath using a four-channel glass pipette,
with the tip placed as close as possible to the clamped cell.
Data acquisition and analysis.
Voltage-clamp commands, data acquisition, and data analysis were
controlled via a computer equipped with a Digidata 1200 interface (Axon
Instruments). pCLAMP software (versions 5.51 and 6.0, Axon Instruments)
was used to generate whole cell current-voltage (I-V) relations, with the membrane currents resulting from voltage stimuli filtered at 1 kHz, sampled at 2.5 kHz, and stored directly on the
computer hard disk. Cells were held at 50 mV, and 400-ms pulses from
100 to +120 mV were applied in 20-mV increments every 2 s.
Cell Volume Measurement
The relative cell volume was monitored by image analysis with
fura 2 as fluorescent volume indicator, as previously reported (22). Six- to 20-day-old cell monolayers grown on petri
dishes were loaded with a solution of 2 µM fura 2 containing 0.01%
pluronic acid for 20 min at 37°C and then washed with an NaCl
solution. The fluorescence was monitored with 360-nm excitation
wavelength. At 360 nm, the variations in the signal emitted by the
probe are directly proportional to the variations in 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 HEPES, pH 7.4 [osmotic pressure
(Posm) = 320 mosmol/kgH2O] at 30 ml/min, and images were averaged eight times and recorded every 5 s for 15 min. Once the fluorescence was stabilized, a hypotonic shock
was induced by perfusing the NaCl solution without mannitol
(Posm = 200 mosmol/kgH2O). The relative
change in cell volume was estimated from the fluorescent signal by
assuming that a 30% decrease in osmolarity caused a decrease in the
fluorescent signal corresponding to a maximum swelling of 30% compared
with the initial volume. The means of relative volume changes were obtained by analysis of 10-20 zones in each culture (n)
chosen with the software. Each zone delimited a cytoplasmic area chosen in individual cells.
Image analysis.
The optical system was composed of a Zeiss ICM-405 inverted microscope
and a Zeiss ×40 objective, which was used for epifluorescent measurement with a 75-W xenon lamp. The excitation beam was filtered through a narrow-band filter centered at 360 nm, mounted in a motorized
wheel (model Lambda 10-2, Sutter Instrument), and 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 eight-bit Extended ISIS camera was equipped with an integration
module to maximize signal-to-noise ratio. The video signal from the
camera proceeded to an image processor integrated in a DT2867 image
card (Data Translation) installed in a Pentium 100 personal computer.
The processor converts the video signal to 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 with the AIW software (version 2.0, Axon Instruments). The
final calculations were made using Excel software (Microsoft).
Calibration.
We used the methods described by Tauc et al. (29) using
2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein and
improved more recently by Raat et al. (17) using fura 2. After cells were loaded with the fluorescent probe in the culture
medium, they were perfused with a solution adjusted to various
osmolarities (150-400 mosmol/kgH2O) by omitting
mannitol. For each osmolarity, two images were stored, averaged, and
subsequently corrected for fading after background subtraction. The
mean fluorescence (360 nm) of five areas was plotted against the
inverse of Posm (in mosmol/kgH2O). Data
showed that when the cells were exposed to a hyposmotic solution,
fluorescence decreased linearly with Posm according to
Boyle's law. To verify that cells in culture behave as osmometers
in a reversible manner, we performed experiments in which the cultures
were perfused successively and randomly with 200-300
mosmol/kgH2O solutions. The fluorescent signal was related
to Posm in a reversible way. In all calibration
experiments, images were recorded 1-2 min after the beginning of
perfusion, at which time the swelling in hypotonic solutions reached
the maximum value. These methods measure variations in the relative volume as a function of Posm of the perfusion medium
(29).
Intracellular Ca2+ Measurements
Intracellular Ca2+ concentration
([Ca2+]i) was measured in cells grown in
petri dishes and loaded for 45 min at room temperature with a solution
of 2 µM fura 2-AM containing 0.01% pluronic acid. The cells were
washed with NaCl solution containing (in mM) 140 NaCl, 5 KCl, 1 MgSO4, 5 glucose, 20 HEPES, pH 7.40, and 1 Tris. Cells were
successively excited at 350 and 380 nm, with images digitized and
stored on the computer hard disk for later analysis. Each raw image was
the result of an integration of four to five frames averaged four
times. The acquisition rate was one image every 10 s. For each
monolayer, [Ca2+]i was monitored in
18-20 random cells. The equation of Grynkiewicz et al.
(9) was used to calculate
[Ca2+]i from the dual
wavelength-to-fluorescence ratio.
Expression in Cultured Cells
The cDNA encoding CFTR was introduced into a polycistronic
expression vector derived from the pIRESneo plasmid (cytomegalovirus promoter; Clontech) in which the neomycin resistance gene had been
replaced by cDNA encoding the chain of the human CD8 cell surface
antigen. Cells were transfected using the DAC-30 method according to
the manufacturer's instructions (Eurogentec, Herstal, Belgium).
Six-day-old cultured cells grown on 35-mm-diameter petri dishes were
serum starved for 24 h before transfection. Transfected cells with
2 µg of CD8-CFTR coexpress CFTR and CD8 at their plasma membrane and
can be visualized using anti-CD8 antibody-coated beads (Dynabeads
M-450, Dynal, Oslo, Norway) (11a). Cells were electrophysiologically tested 48 h after transfection.
Chemicals
5-Nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB; Calbiochem)
was prepared at 100 mM in DMSO and used at 0.1 mM in final solutions.
4-4'-Diisothiocyanostilbene-2,2'-disulfonic acid (DIDS) was
directly dissolved at a final concentration of 1 mM. Forskolin and
ionomycin were prepared at 10 and 2 mM, respectively, in ethanol and
used at 10 and 2 µM, respectively, in bath medium. DIDS, forskolin, ARL-67156
(6-N,N-diethyl-
-
-dibromomethylene-D-adenosine-5'-triphosphate trisodium), apamin, and ionomycin were obtained from Sigma (Saint Quentin Fallavier, France). Fura 2-AM (Molecular Probes) was dissolved at 3 mM in DMSO and added to the loading solution at a final
concentration of 2 µM, along with 0.01% pluronic acid.
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RESULTS |
Cl Currents Activated by Forskolin
Experiments were performed in a hyperosmotic extracellular
solution (350 mosmol/kgH2O) to characterize
Cl currents activated by forskolin in PCT, DCT, and CCT
cells. Under these conditions, volume-activated Cl
currents could not be detected. In the absence of forskolin, the
voltage-step protocol elicited small currents that changed linearly
with the membrane potential in PCT, DCT, and CCT cell cultures from
kidneys of cftr+/+ and cftr/ mice
(data not shown). In cftr+/+ mice, exposure of cultured DCT
and CCT cells to 10 µM forskolin induced an increase in membrane
current amplitudes (Fig. 1A)
that reached a peak value 3-4 min after the beginning of the
perfusion. These activated currents exhibited a linear I-V
relationship, with a reversal potential (Erev)
of 1.3 ± 2.3 mV and a conductance of 6.4 ± 0.6 nS in DCT
cells and an Erev of 2.2 ± 2.9 mV and a
conductance of 6.0 ± 1.2 nS in CCT cells (n = 8 monolayers from 4 mice). In contrast, application of forskolin did not
modify the currents recorded in cultured PCT cells (Fig. 1A). The unstimulated whole cell current in these cells
reversed at 3.1 ± 1.2 mV, with a slope conductance of 1.1 ± 0.1 nS (n = 9 monolayers from 4 mice). As
expected, in cftr/ mice, addition of
forskolin did not stimulate Cl conductance in all the
cultured segments studied. This observation clearly indicated that, in
DCT and CCT cells from cftr+/+ mice, the Cl
conductance stimulated by forskolin was related to CFTR.
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Fig. 1.
A: forskolin-induced whole cell Cl
currents in proximal convoluted tubule (PCT), distal convoluted tubule
(DCT), and cortical collecting tubule (CCT) cells in primary culture of
cftr+/+ and cftr/ mice. Membrane
voltage was held at 50 mV and stepped to test potential of 100 to
+120 mV in 20-mV increments. Whole cell currents were recorded after 3 min of extracellular perfusion of 10 µM forskolin in the presence of
1 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2
in extracellular bath. B: effects of 1 mM DIDS, 0.1 mM
5-nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB), and
extracellular Cl substitution by I on
forskolin-induced whole cell Cl currents measured at +100
mV. Values are means ± SE; n, number of monolayers
from 4 different mice.
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The forskolin-sensitive Cl currents measured at +100 mV
are compared in primary cultures of PCT, DCT, and CCT from
cftr+/+ and cftr/ mice in Fig.
1B. Only DCT and CCT from wild-type mice exhibited
forskolin-activated Cl currents that were blocked by 0.1 mM NPPB and insensitive to 1 mM DIDS in the extracellular bath.
Moreover, in these segments, replacing external Cl with
I strongly inhibited the Cl currents
activated by forskolin and caused Erev to shift
toward positive values: Erev for
I = 37.5 ± 1.4 and 17.0 ± 6.8 mV for DCT
and CCT, respectively (n = 4 monolayers from 4 different mice).
Ca2+-Induced
Cl Currents
Whole cell currents were recorded with Ca2+-free (1 mM
EGTA) solutions containing NMDG chloride as the major cation in the
pipette and with extracellular solutions containing NMDG chloride and 1 mM CaCl2. The extracellular solution was adjusted to 350 mosmol/kgH2O with mannitol to avoid inducing
volume-activated currents. The control macroscopic currents were
recorded, and 2 µM ionomycin was added to the NMDG chloride bathing
solution. Stimulated currents were recorded after 2 min. Figure
2A shows the currents recorded in PCT, DCT, and CCT monolayers from cftr+/+ and
cftr/ mice. In all cultured segments from
both types of mice, addition of ionomycin stimulated Cl
currents, which increased during depolarizing voltage pulses. The
kinetics of the macroscopic current were clearly time dependent for
depolarizing potentials with a slowly developing component. In cultured
PCT cells from cftr+/+ mice, currents reversed at 2.6 ± 0.3 mV (n = 16 monolayers from 4 different mice).
Instantaneous currents measured 5 ms after the beginning of the
stimulation were almost linear, with an inward current of 485 ± 34 pA at 100 mV and an outward current of 635 ± 42 pA at +100
mV. The steady-state current at 380 ms exhibited a marked outward
rectification, with an inward current of 352 ± 40 pA at 100 mV
and an outward current of 834 ± 79 pA at +100 mV
(n = 16 monolayers from 4 different mice). When the
steady-state current measurements were used to calculate the
Cl conductance, the maximal outward conductance was
significantly different from the maximal inward conductance: 9.6 ± 0.8 and 2.4 ± 0.4 nS, respectively (n = 16 monolayers from 4 different mice; P < 0.02).
Finally, as shown in Fig. 2B, 1 mM DIDS inhibited the ionomycin conductance by 85.6 ± 5.1% (n = 16 monolayers from 4 different mice). In cultured DCT and CCT cells from
cftr+/+ mice, the Cl currents induced by
ionomycin strongly resembled those induced in cultured PCT cells: the
steady-state currents were outwardly rectifying and strongly blocked by
1 mM DIDS (Fig. 2B). Moreover, as illustrated in Fig. 2, the
ionomycin-sensitive Cl conductances measured in
cftr/ mice exhibited characteristics roughly
similar to those measured in cftr+/+ mice, indicating that
CFTR does not participate in the Ca2+-sensitive
Cl conductance along the mouse nephron.
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Fig. 2.
A: Ca2+-induced whole cell Cl
currents in PCT, DCT, and CCT cells in primary culture of
cftr+/+ and cftr/ mice. Membrane
voltage was held at 50 mV and stepped to test potential of 100 to
+120 mV in 20-mV increments. Whole cell currents were recorded after 2 min of extracellular perfusion of 2 µM ionomycin in the presence of 1 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2
in extracellular bath. B: effects of 1 mM DIDS on
Ca2+-induced whole cell Cl currents.
Steady-state currents at +100 mV were measured 380 ms after onset of
pulse. Values are means ± SE; n, number of monolayers
from 4 different mice.
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Cl Currents Induced by a Hypotonic
Shock
To study the effects of changes in Posm on the
development of Cl conductance, currents were induced by
osmotic shock. Whole cell currents were recorded with
Ca2+-free (5 mM EGTA) pipette solutions containing NMDG
chloride and maintained at 290 mosmol/kgH2O. Moreover, to
eliminate any participation of cations in the inward current,
experiments were carried out after Na+ in the bath solution
was replaced with NMDG chloride and in the presence of 1 mM
CaCl2. In cftr+/+ mice, the control currents were first measured in cultured PCT, DCT, and CCT cells with an extracellular solution osmolarity of 350 mosmol/kgH2O.
Under this condition, the voltage-step protocol elicited small
time-independent currents that changed linearly with the membrane
voltage and had Erev of 6.2 ± 1.5, 1.5 ± 1.6, and +1.9 ± 0.8 mV for PCT, DCT, and CCT cells,
respectively (n = 4 monolayers from 4 different mice). Because of their small amplitude, the nature of these currents was not analyzed further.
The monolayers were then perfused with a 290 mosmol/kgH2O
solution. Figure 3A gives the
currents recorded in PCT, DCT, and CCT cells. In >95% of the
cftr+/+ cells, an increase in the whole cell current was
observed within 1 min. In all epithelial cell types, the currents
reached a maximum after 4-5 min. Under these conditions, the
initial currents recorded at +100 mV were ~2.5 times the amplitude of
the currents recorded at 100 mV. These large, outwardly rectifying
currents showed a small time-dependent inactivation at depolarizing
potentials 
60 mV in cultured PCT and CCT cells and 40 mV in
cultured DCT cells. In most cases, the time course of this inactivation
could be well fitted with a single exponential irrespective of the
recording time. When the cells were reexposed to the hyperosmotic
solution, the currents returned to the control level within 2-3
min (Fig. 3B). In the three cultured segments, the
currents induced by hypotonicity were strongly blocked by 1 mM DIDS
(Fig. 3B).
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Fig. 3.
A: characteristics of swelling-induced whole cell
Cl currents in PCT, DCT, and CCT cells in primary culture
of cftr+/+ and cftr/ mice.
Membrane voltage was held at 50 mV and stepped to test potential of
100 to +120 mV in 20-mV increments. Whole cell currents were recorded
after 4-5 min of extracellular perfusion of a 30% hypotonic
solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette
solution and 1 mM CaCl2 in extracellular bath.
B: effects of 1 mM DIDS and hyperosmotic solution (350 mosmol/kgH2O) on swelling-induced whole cell
Cl currents. Steady-state currents at +100 mV were
measured 20 ms after onset of pulse. Values are means ± SE;
n, number of monolayers from 4 different mice.
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In cftr/ mice, hypotonic shock was completely
inefficient for increasing Cl conductance in the
three different cultured segments (Fig. 3, A and
B). In all nephron segments studied, an absence of response to hypotonic shock was observed in 100% of the recorded cells. This
result implicates CFTR in the control of the swelling-activated Cl conductance in renal epithelium.
The results reported above clearly show that Cl
conductances developed in the presence of forskolin, Ca2+,
or hypotonic shock in DCT cells were roughly similar to those recorded
in CCT cells under the same experimental conditions. Therefore, in the
following experimental series, no distinction was made between DCT and
CCT cells.
Cl Currents in Cultured PCT and DCT
Cells From cftr/
Mice Transfected With CFTR cDNA
The cftr/ PCT and DCT cells in primary
culture were transfected with CD8-CFTR plasmid, which allows
visualization of transfected cells using anti-CD8 antibody-coated
beads. After 48 h of transfection, whole cell currents of coated
cells were recorded and compared with whole cell currents of control
unlabeled cells. Figures 4 and
5 illustrate the currents recorded in
cftr/-transfected PCT and DCT cells in the
presence of 10 µM forskolin. As expected, addition of forskolin
induced an increase in membrane current amplitudes in PCT (Fig.
4Ab) and DCT (Fig. 5Ab) cells coated with beads
only. Figures 4Ae and 5Ae show that the
forskolin-activated currents exhibited a linear I-V
relation, with an Erev of 0.2 ± 0.1 mV and
a conductance of 7.6 ± 0.4 nS for PCT cells (n = 5 cells) and an Erev of 0.16 ± 0.5 mV and
a conductance of 8.3 ± 0.5 nS for DCT cells (n = 7 cells). Currents in both cell types were insensitive to 1 mM DIDS
(Figs. 4Ac and 5Ac) and blocked by 77 ± 3 and 85 ± 2% for PCT and DCT, respectively, when Cl
was replaced by I (Figs. 4Ad and
5Ad). Moreover, this substitution shifted the Erev toward the more positive value:
Erev for I = 36.4 ± 8 and 25.7 ± 9 mV in PCT and DCT, respectively. Overall, these
forskolin-sensitive Cl currents were identical to those
measured in cftr+/+ DCT cells, indicating that transfection
with CFTR plasmid could restore the normal CFTR currents in PCT and DCT
cftr/ cells.
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Fig. 4.
Restoration of CFTR currents and swelling-activated
Cl currents by transitory transfection of
pIRES-CD8-cftr in PCT cells from
cftr/ mice. Transfected cells were visualized
using anti-CD8 antibody-coated beads. Membrane potential was held at
50 mV and stepped to test potential of 100 to +120 mV in 20-mV
increments. A: CFTR currents in cells labeled with
anti-CD8-coated beads. a: Control; b: 10 µM
forskolin in bath solution; c: 10 µM forskolin + 1 mM
DIDS; d: 10 µM forskolin with extracellular substitution
of Cl by I. e: Average
current-voltage (I-V) relationships measured 200 ms after
onset of pulse, obtained from the same cell at rest, during forskolin
stimulation alone and after Cl substitution by
I. Values are means ± SE of 5 cells from 3 transfected monolayers. B: swelling-activated
Cl currents recorded after 4-5 min of extracellular
perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and
5 mM MgATP in pipette solution and 1 mM CaCl2 in
extracellular bath. a-c: Whole cell currents in cells
labeled with anti-CD8-coated beads. d: Average
I-V relationships measured 20 ms after onset of pulse,
obtained from the same cell at rest (B), during perfusion
with hyposmotic solution, and after perfusion with hyperosmotic
solution. Values are means ± SE of 4 cells obtained from 3 transfected monolayers.
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Fig. 5.
Restoration of CFTR currents and swelling-activated
Cl currents by transitory transfection of
pIRES-CD8-cftr in DCT cells from
cftr/ mice. Transfected cells were visualized
using anti-CD8 antibody-coated beads. Membrane potential was held at
50 mV and stepped to test potential of 100 to +120 mV in 20-mV
increments. A: CFTR currents in cells labeled with
anti-CD8-coated beads. a: Control; b: 10 µM
forskolin in bath solution; c: 10 µM forskolin + 1 mM
DIDS; d: 10 µM forskolin with extracellular substitution
of Cl by I. e: Average
I-V relations measured 200 ms after onset of pulse, obtained
from the same cell at rest, during forskolin stimulation alone and
after Cl substitution by I. Values are
means ± SE of 7 cells from 3 transfected monolayers. B
and C: swelling-activated Cl currents recorded
after 4-5 min of extracellular perfusion of a 30% hypotonic
solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette
solution and 1 mM CaCl2 in extracellular bath. Ba,
Bb, and Bc: whole cell currents in cells labeled with
anti-CD8-coated beads. Bd: average I-V
relationships measured 20 ms after onset of pulse, obtained from the
same cell at rest (B), during perfusion with hyposmotic
solution, and after perfusion with hyperosmotic solution. Values are
means ± SE of 4 cells from 3 transfected monolayers.
C: whole cell currents in cells not labeled with
anti-CD8-coated beads.
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In another experimental series, the effect of a hypotonic shock was
studied in cftr/ PCT and DCT cells
transfected with the cftr plasmid. In both cell types, after
the hypotonic shock, the coated cells developed Cl
currents within 3 min (Figs. 4B and 5B). The
initial currents measured 20 ms after the onset of the voltage pulse
rectified in the outward direction (Figs. 4Bd and
5Bd). For PCT cells, they reversed at +0.6 ± 0.4 mV
(n = 4 cells), and the total current at +100 mV was 3.8 times that at 100 mV: 1,555 ± 150 vs. 405 ± 18 pA
(n = 4 cells). For DCT cells, they reversed at
+0.9 ± 0.3 mV (n = 4 cells), and the total
current at +100 mV was 2.2 times that at 100 mV: 1,123 ± 155 vs. 508 ± 86 pA (n = 4 cells). These large
outwardly rectifying currents showed time-dependent inactivation at
depolarizing step potential >40 mV. Finally, replacement of the
hypotonic bath solution by a hypertonic solution inhibited the
Cl currents by 74 ± 3 and 80 ± 4% for PCT
and DCT cells, respectively (n = 4). As expected, the
uncoated cells remained insensitive to the hypotonic shock (Fig.
5B). Therefore, transfection of CFTR also restores the
swelling-activated Cl conductance in
cftr/ PCT and DCT cells.
Regulation of the Cl Conductance
Induced by Hypotonic Shock in
cftr+/+ and
cftr/ DCT and CCT Cells
Role of extracellular Ca2+ in the
presence of high EGTA concentration in the pipette solution.
In cftr+/+ cells, 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 EGTA without additional
Ca2+. The effects of extracellular Ca2+ on the
development of hypotonicity-induced Cl currents were also
tested in cftr+/+ DCT and CCT cells. When the hypotonic
shock was carried out in the absence of bath Ca2+,
development of the Cl current was significantly impaired
(Fig. 6A). As previously
reported in rabbit distal bright convoluted tubule (DCTb) in primary
culture (20), these experiments confirm that extracellular
Ca2+ was required to activate the swelling-activated
Cl conductance in DCT and CCT cells cultured from
cftr+/+ mice. Using this information, we therefore decided
to study the effect of an influx of Ca2+ on
swelling-activated Cl conductance in
cftr/ DCT and CCT cells. For this purpose,
the effects of ionomycin were tested on whole cell Cl
currents recorded in the absence of intracellular free
Ca2+. Whole cell currents were recorded in the presence of
20 mM EGTA in the pipette solution and 1 mM free Ca2+ in
the bath (Fig. 6B). In the absence of ionomycin in the bath solution, the hypotonic shock remained inefficient for triggering Cl currents in cftr/ cells
(Fig. 6Ba). In contrast, when the hypotonic shock was
performed in the presence of 2 µM ionomycin, Cl
currents were activated within 5 min (Fig. 6Bb). These
currents showed time-dependent inactivation at depolarizing step
potentials >60 mV and displayed an outwardly rectified instantaneous
I-V plot (Fig. 6Be) with an
Erev of + 1.1 ± 0.3 mV
(n = 7). When the cells were reexposed to the
hyperosmotic solution, the currents returned toward control level
within 2-3 min (Fig. 6, Bc and Be). Alternatively, addition of DIDS rapidly reduced the Cl
currents (89.7 ± 4% inhibition at +100 mV, n = 5; Fig. 6Bd). Overall, the ionomycin-induced
Cl currents developed during hypotonicity in DCT and CCT
cells from cftr/ mice were quite similar to
the swelling-activated Cl currents measured in
cftr+/+ mice.
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Fig. 6.
A: effect of extracellular Ca2+ on
development of hypotonicity-induced Cl currents in
cultured DCT cells from cftr+/+ mice. Membrane voltage was
held at 50 mV and stepped to test potential of 100 to +120 mV in
20-mV increments. Whole cell currents were recorded after 4-5 min
of extracellular perfusion of a 30% hypotonic solution in the presence
of 5 mM EGTA in pipette solution and in the absence of extracellular
Ca2+ in bath solution. B: effects of ionomycin
on development of Cl currents in cultured DCT cells from
cftr/ mice. Membrane voltage was held at 50 mV and
stepped to test potential of 100 to +120 mV in 20-mV increments.
Whole cell currents were recorded after 4-5 min of extracellular
perfusion of a 30% hypotonic solution in the presence of 20 mM EGTA
and 5 mM MgATP in pipette solution and 1 mM CaCl2 in
extracellular bath. Whole cell currents were measured during hypotonic
shock. a: Control cells; b: 2 µM ionomycin;
c: 2 min of replacement of extracellular solution with a
hypertonic solution; d: 1 mM DIDS. e: Average
I-V relationships measured 20 ms after onset of pulse,
obtained from the same cell at rest. Values are means ± SE;
n, number of cells from 4 monolayers.
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Very similar results were obtained with PCT cells in primary culture.
Briefly, in cftr+/+ cells, the swelling-sensitive
Cl conductance depended on external Ca2+, and
in cftr/ cells, this conductance could be
reactivated by addition of ionomycin to the bath solution (data not shown).
Role of extracellular Ca2+ in the
absence of EGTA in the pipette solution.
The experiments described above indicate that Ca2+ influx
induced by ionomycin could restore the swelling-activated
Cl currents in cftr/ cells. To
further analyze this phenomenon, the effect of ionomycin was tested in
the absence of EGTA in the pipette solution. Two successive, increased
external Ca2+ concentrations were applied to the same
cftr/ DCT cells. The results are reported in
Fig. 7A. Control currents were
recorded, and the cells were perfused with a Ca2+-free
solution containing 2 µM ionomycin. After 2 min, raising the
Ca2+ concentration to 0.1 µM induced Cl
currents that were identical to the swelling-activated Cl
currents (Fig. 7Ab). A further new increase in
Ca2+ concentration to 1 µM enhanced the currents (Fig.
7Ac). These currents showed virtually no inactivation during
the 400-ms voltage pulse. Currents obtained by subtracting the current
recorded at 0.1 µM external Ca2+ from that recorded at 1 µM Ca2+ are shown in Fig. 7Ad. The resulting
currents exhibited the characteristic profile of the
Ca2+-sensitive Cl currents.
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Fig. 7.
A: effect of extracellular Ca2+
concentration ([Ca2+]ext) on development of
Cl currents in cultured DCT cells from cftr+/+
mice. Whole cell currents were recorded in the absence of EGTA in
pipette solution. a: Control; b: 2 min of
extracellular perfusion of 0.1 µM Ca2+ in the presence of
ionomycin; c: 2 min of extracellular perfusion of 1 µM
Ca2+ in the presence of ionomycin; d: currents
obtained by subtraction of c from b using
pCLAMPFIT 6.0 software. e: Average I-V
relationships measured 20 ms after onset of pulse, obtained from the
same cell at rest, during perfusion of ionomycin in the presence of
different Ca2+ concentrations. Values are means ± SE
of 6 cells obtained from 6 monolayers. B: effect of
hypotonic shock on intracellular free Ca2+ concentration
([Ca2+]i) in fura 2-loaded DCT cells from
cftr+/+ and cftr/ mice. Fura 2 fluorescence
was monitored and converted to [Ca2+]i
as described in MATERIALS AND METHODS. Hypotonic shock was
induced by perfusion of a hypotonic NaCl solution (200 mosmol/kgH2O). Values are means ± SE of 20 random
cells from 3 monolayers.
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On the basis of these results, it appears that Ca2+ entry
is an important step in the development of swelling-activated
Cl conductance. Fluorescence experiments using fura
2-loaded DCT cells were therefore carried out to follow cytosolic
Ca2+ variations during hypotonic shock. The effect of
hypotonic solution on [Ca2+]i in DCT cells
from cftr+/+ and cftr/ mice is
shown in Fig. 7B. In both types of mice, when the cells were
bathed with an isotonic NaCl solution (300 mosmol/kgH2O)
containing 1 mM CaCl2, the resting
[Ca2+]i averaged 30.8 ± 5.2 nM
(n = 20). Swelling the cftr+/+ DCT cells with hypotonic NaCl solution (200 mosmol/kgH2O) induced a
transient increase of [Ca2+]i that reached a
maximum value of 120.1 ± 25.1 nM and returned close to the
control value within 5 min (Fig. 7B). In contrast, swelling
the cftr/ DCT cells did not significantly
modify [Ca2+]i.
Role of extracellular adenosine.
We previously demonstrated that stimulation of A1 adenosine
receptors could be implicated in the control of swelling-induced Cl currents in rabbit DCT (20), and
experiments were therefore performed to determine the role of adenosine
in Cl permeability of PCT and DCT cells from
cftr+/+ and cftr/ mice. Results of
whole cell experiments performed in cftr/ PCT
and DCT cells are illustrated in Fig. 8.
These results were strictly identical to those obtained with
cftr+/+ PCT and DCT cells. In both types of primary
cultures, 10 µM adenosine activated an outwardly rectifying
Cl conductance with a time-dependent inactivation at
depolarizing potentials and with a maximal effect at 3-4 min (Fig.
8A). Erev of the stimulated current
were 3.8 ± 3.7 mV (n = 5 monolayers) and 0.3 ± 2.9 mV (n = 4) for PCT and DCT cells, respectively. In the presence of adenosine, the maximal slope conductances reached 19 ± 9 nS (n = 5) and 11 ± 4 nS
(n = 4) in PCT and DCT cells, respectively. These
adenosine-sensitive Cl currents were decreased in the
presence of 1 mM DIDS by 90 and 78% in PCT and DCT cells,
respectively. To determine whether the response to adenosine occurred
via receptor-mediated mechanisms, we examined the effect of a
P1-selective receptor antagonist, 8-cyclopentyl-1,3-diproxylzanthine (DPCPX). Treatment of DCT cells with
10 µM DPCPX completely inhibited the development of outward Cl currents first induced by 10 µM adenosine in PCT and
DCT cells (Fig. 8).
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Fig. 8.
Effects of adenosine on development of Cl currents in
cultured PCT (A) and DCT (B) cells from
cftr/ mice. Membrane voltage was held at 50 mV and
stepped to test potential of 100 to +120 mV in 20-mV increments.
Whole cell currents were recorded after 4-5 min of extracellular
perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and
5 mM MgATP in pipette solution and 1 mM CaCl2 + 10 µM adenosine in extracellular bath. DIDS (1 mM) was perfused
after development of Cl currents. Cells were treated with
antagonist 8-cyclopentyl-1,3-diproxylxanthine (DPCPX) before
exposure to adenosine.
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The effect of adenosine was concentration dependent. The dose-response
curve in cultured DCT cells from cftr+/+ mice is shown in
Fig. 9. The half-maximal effect from this
curve occurred at 5.0 × 107 M adenosine and the
maximal effect at >105 M.
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Fig. 9.
Adenosine dose-response curve. Membrane voltage was held
at 50 mV and stepped to test potential of 100 to +120 mV in 20-mV
increments. Whole cell currents were recorded after 3 min of
extracellular perfusion of adenosine at different concentrations in
isotonic N-methyl-D-glucamine solutions in the
presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM
CaCl2 in extracellular bath. Values at +100 mV were
converted to percent activation. Values are means ± SE of 6 cells
from 3 monolayers.
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Role of extracellular ATP.
In addition to adenosine, it has been postulated that ATP could
activate a volume-sensitive-like Cl conductance in
immortalized rabbit distal cells (20). To check this
possibility in PCT and DCT cells from cftr+/+ and
cftr/ mice, we studied the role of ATP in the
control of whole cell Cl currents in the presence of 5 mM
EGTA in the pipette solution. In PCT and DCT monolayers, addition of 10 µM ATP to the bath solution induced activation of Cl
currents within 4-5 min. This ATP-activated Cl
current showed time-dependent inactivation at depolarizing step potentials >60 mV (Fig. 10,
A and B) and displayed an outwardly rectified
instantaneous I-V plot (data not given) with
Erev close to 0 mV. DIDS (1 mM) strongly
decreased ATP-activated currents in both types of monolayers. Overall,
these currents were quite similar to those induced by adenosine.
Moreover, the effect of ATP was completely blocked by 10 µM DPCPX,
indicating that the action was triggered via P1, rather
than P2, receptors. Such results suggested that stimulation
of Cl currents in the presence of ATP was most probably
due to an action of adenosine generated by degradation of ATP.
Experiments were therefore carried out to check this hypothesis. For
this purpose, DCT cells from cftr+/+ mice were subjected to
a hypotonic shock in the presence of the selective ecto-ATPase
inhibitor ARL-67156. ARL-67156 (100 µM) completely blocked the
swelling-activated Cl currents (Fig. 10C).
Adenosine (10 µM) restored a swelling-activated Cl
conductance, which displayed an outwardly rectified instantaneous I-V plot with Erev close to 0 mV
(Fig. 10Cd) and was strongly inhibited by DIDS (Fig.
10Cb).
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Fig. 10.
Effects of ATP on development of Cl currents in
cultured PCT (A) and DCT (B) cells from
cftr/ mice. Membrane voltage was held at 50 mV and
stepped to test potential of 100 to +120 mV in 20-mV increments.
Whole cell currents were recorded after 4-5 min of extracellular
perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and
5 mM MgATP in pipette solution and 1 mM CaCl2 + 10 µM ATP in extracellular bath. DIDS (1 mM) was perfused after
development of Cl currents. Cells were treated with DPCPX
before exposure to adenosine. Values are means ± SE;
n, number of cells from 3 monolayers. C: effects
of ecto-ATPase antagonist ARL-67156 on swelling-activated
Cl currents in DCT cells from cftr+/+ mice.
Whole cell currents were recorded after 4-5 min of extracellular
perfusion of a hypotonic solution in the presence of 100 µM ARL-67156
(a), 10 µM adenosine (b), or 1 mM DIDS
(c). d: Average I-V relationships
measured 20 ms after onset of pulse obtained from the same cell at
rest. Values are means ± SE of 5 cells from 3 monolayers.
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RVD in PCT and DCT Cells From
cftr+/+ and
cftr/ Mice
To confirm the role of swelling-activated Cl
currents in cell volume regulation, we measured relative cell volume in
monolayers by fluorescence-video microscopy in PCT and DCT cells from
cftr+/+ and cftr/ mice. As
expected, PCT and DCT cells swelled in response to a hypotonic shock
(Fig. 11). This cell swelling was
followed by an RVD in PCT and DCT cells from cftr+/+ mice.
At 2 min after the hypotonic shock, the relative cell volume reached
130 ± 2% (n = 3 monolayers) and 120 ± 1%
(n = 8 monolayers) of the initial volume in PCT and DCT
cells, respectively. PCT cells returned to 105 ± 1% of their
original volume within 4 min (Fig. 11A), whereas DCT cells
recovered 104 ± 1% of their volume within only 2 min (Fig.
11B). The RVD phenomenon in both cell types was inhibited in
the presence of 100 µM NPPB (Fig. 11). Contrary to observations in
cells from cftr+/+ mice, the RVD mechanism was completely
impaired in PCT and DCT cells from cftr/ mice. When
perfused with the hypotonic solutions, these cells never returned to
their initial volume. Addition of 10 µM adenosine during hypotonic
shock restored RVD in DCT cells from cftr/ mice (Fig.
11B).
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Fig. 11.
Effects of hypotonic shock on cell volume in PCT and DCT
cells from cftr+/+ and cftr/ mice.
Cultures were loaded with 2 µM fura 2 and rinsed in an isotonic NaCl
solution (300 mosmol/kgH2O) for 3 min. A hypotonic shock
was induced by reducing osmolarity of NaCl solution to 200 mosmol/kgH2O. Images were recorded every 15 s. After
analysis, relative volume change as percentage of initial volume was
plotted against time. A: regulatory volume decrease (RVD) in
3 monolayers from PCT cells (25 random cells each) in the presence or
absence of 100 µM 5-nitro-2-(3-phenylpropylamino)-benzoic acid
(NPPB). B: RVD in 10 monolayers from DCT cells (25 random
cells each) from cftr/ mice and 8 monolayers (25 random
cells each) from DCT cells from cftr+/+ mice in the presence
or absence of 100 µM NPPB or adenosine.
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DISCUSSION |
The aim of the present study was to investigate the putative role
of CFTR in the control of Cl conductances along the
different segments of the mouse nephron. Using the patch-clamp
technique to measure whole cell conductance, we analyzed three distinct
types of Cl currents in primary cultures of PCT, DCT, and
CCT segments obtained by microdissection of kidney cortex from
wild-type cftr+/+ and cftr/ mice. These
Cl conductances consisted of forskolin-activated,
volume-sensitive, and Ca2+-activated Cl currents.
In the first series of experiments, the effect of forskolin on
Cl conductance was tested in PCT, DCT, and CCT cells. For
this purpose, swelling-activated currents were blocked by exposing the
cells to a hyperosmotic solution, and Ca2+-activated
conductances were impaired by the use of high EGTA concentrations in
the pipette solution. In cftr+/+ mice, external application
of forskolin activated a linear Cl current in DCT and
CCT, but not PCT, cells. The halide selectivity was consistent with low
relative I permeability and with an inhibitory effect of
I. Moreover, this forskolin-stimulated conductance was
blocked by NPPB but was quite insensitive to DIDS. These
characteristics are very similar to those reported previously in rabbit
distal bright convoluted tubule (DCTb) in primary culture
(21). In contrast, in cultured DCT and CCT cells from
cftr/ mice, addition of forskolin remained completely
inefficient for increasing Cl conductances. Taken
together, the results obtained in primary cultures from
cftr+/+ and cftr/ mice clearly demonstrate
that, at least in DCT and CCT cells, the activity of
forskolin-activated Cl channels is consistent with CFTR.
In other words, as we concluded in a previous study (21),
the channel involved in the Cl currents activated by
forskolin in DCT and CCT is the small-conductance CFTR Cl channel.
Interestingly, application of forskolin did not stimulate any
Cl current in primary cultures of PCT cells from
cftr+/+ mice. Such an observation was reported in primary
culture of rabbit PCT, in which no CFTR expression and no
forskolin-activated Cl currents were detected in the
apical membrane, despite the presence of CFTR mRNA (21).
The presence of CFTR in the mammalian kidney is now well documented
(2, 15, 16), but the absence of detectable renal disease
in CF patients led several authors to postulate that an increase of
another type of Cl channel might compensate for the lack
of cAMP-activated Cl channels in renal tissue (6,
11). On the other hand, the cftr/ mice
used in the present study did not present significant pulmonary
disease. Moreover, in these mice, it has been shown that the
Ca2+-activated Cl channels could be
candidates for compensation of the missing CFTR Cl
channels (6). To determine whether this possibility could arise in the renal epithelium, we studied the
Ca2+-activated conductance in PCT, DCT, and CCT cells. As
expected, in cftr+/+ mice, extracellular application of
ionomycin rapidly activated currents in all types of monolayers. This
Ca2+-sensitive conductance was similar to that previously
described in rabbit PCT and DCTb cells under identical experimental
conditions (21, 22). In cftr/ mice, the
increase of Cl conductance triggered by ionomycin was
strikingly identical to that observed in wild-type mice, eliminating
the hypothesis that Ca2+-activated conductance could
substitute for CFTR Cl conductance in renal epithelium.
In cftr+/+ mice, cultured PCT, DCT, and CCT cells developed
a volume-sensitive Cl current when exposed to a hypotonic
shock. The biophysical and pharmacological characteristics of this
Cl conductance show strong similarities to the properties
of swelling-activated Cl currents described in many other
epithelial cells, including rabbit PCT and DCTb in primary culture
(7, 25). Null mutation of the cftr gene
strongly impaired the swelling-activated Cl currents in
the three different nephron segments. Moreover, cftr/ DCT or CCT cells transfected with cftr cDNA displayed
complete restoration of cAMP-dependent and swelling-activated
Cl currents. Transfection of PCT cells with cftr
cDNA also restored both conductances. These observations indicate
that PCT cells have maintained their ability to insert exogenous CFTR
into the apical membrane. Therefore, the lack of forskolin-induced
Cl conductance in wild-type PCT cells is probably due to
a difference in the protein function, rather than a modification of the
intracellular trafficking leading to protein retention in intracellular membranes.
It is well established that the swelling-activated Cl
channels participate in the RVD phenomenon, which is induced by
exposure of cells to hyposmotic solutions. In the present study, to
determine whether cultured PCT, DCT, and CCT cells develop RVD after a
hypotonic shock, we used a simple fluorescence method for studying
relative cell volume variations (29). The findings
indicate that cultured cells from cftr+/+ mice are sensitive
to osmolarity changes in the bathing medium and that they are capable
of RVD after hypotonic shock. RVD was also examined in cultured cells
from cftr/ mice. These cells exhibited a defective
volume regulation after a hyposmotic shock. This observation confirms
the results in the literature (31) and indicates that CFTR
could play a role in the RVD of epithelia. Obviously, this defective
RVD is due to the fact that the hypotonic shock is completely
inefficient for increasing Cl conductances. The
intervention of CFTR in the control of swelling-activated Cl conductances has been proposed by Chan et al.
(5), who demonstrated, in the human colonic cell line T84,
that an antibody against CFTR inhibited the cAMP- as well as the
swelling-induced whole cell Cl conductances but did not
affect the Ca2+-activated Cl channel. In
previous studies, we found that development of Cl
conductance after a hypotonic shock in rabbit DCT cells was related to
an influx of external Ca2+ through Ca2+
channels (21, 22). Such a hypothesis could also apply to the data obtained in DCT cells from cftr+/+ mice, because
removal of external Ca2+ just before the hypotonic shock
completely impaired the increase in Cl current,
suggesting that Ca2+ influx could participate in activation
of the Cl channels in mouse kidney. It is now proposed
that CFTR could regulate other ion channel proteins (23)
and could also be implicated in different cell functions such as
apoptosis (1, 8, 13) or cytosolic Ca2+
regulation (19). Moreover, a recent study by Braunstein et al. (3) clearly demonstrated that CFTR participates in
cell volume regulation via control of ATP release. Taken together, these observations led us to propose the hypothesis that the absence of
swelling-activated Cl conductance in DCT cells from
cftr/ could be related to an alteration of the
Ca2+ entry. This hypothesis is strengthened by two main
results: 1) The hypotonic shock induced an increase of
[Ca2]i in cftr+/+ DCT cells, but
not in cftr/ cells. 2) In
cftr/ cells, addition of ionomycin in the presence of
high intracellular EGTA concentration restored the ability of the cells
to respond to the hypotonic shock by increasing swelling-sensitive
Cl conductance. It remains to be shown how intracellular
Ca2+ can increase in the presence of a high EGTA
concentration. The observations of Evans and Marty (6a) 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 in the presence of EGTA. In accordance with the model proposed by
Braunstein et al. (3), the defect in cell volume regulation that we observed in cftr/ renal cells could
be due to a defect in the ATP release pathway. We have proposed
(20) that hypotonic shock stimulates ATP release from
rabbit DCT cells. A1 receptors are then activated by
adenosine generated by the degradation of ATP by membrane ectoenzymes,
and this stimulation of A1 receptors induces an influx of
extracellular Ca2+. Finally, this Ca2+ influx
activates the Cl channel. The observation that adenosine
restores swelling-activated Cl conductance and RVD in
cftr/ cells confirms that adenosine is a mediator of
RVD, at least in renal epithelium. Therefore, in cftr/
cells, there is no volume-sensitive ATP release, and the cascade of
events that triggers the final increase in Cl conductance
no longer occurs.
The present results confirm that autocrine ATP rele
currents in primary cultures of mouse
nephron
TOP
ABSTRACT
INTRODUCTION
