1 Unitat de Senyalització Cellular, Departament de Ciències Experimentals i de la Salut, Universitat Pompeu Fabra, 08003 Barcelona, Spain; and 2 Medical Research Council Clinical Sciences Centre, Imperial College, London W12 0NN, United Kingdom
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ABSTRACT |
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The cell
regulatory volume decrease (RVD) response triggered by hypotonic
solutions is mainly achieved by the coordinated activity of
Cl and K+ channels. We now describe the
molecular nature of the K+ channels involved in the RVD
response of the human bronchial epithelial (HBE) cell line 16HBE14o
.
These cells, under isotonic conditions, present a K+
current consistent with the activity of maxi K+ channels,
confirmed by RT-PCR and Western blot. Single-channel and whole cell
maxi K+ currents were readily and reversibly activated
following the exposure of HBE cells to a 28% hypotonic solution. Both
maxi K+ current activation and RVD response showed calcium
dependency, inhibition by TEA, Ba2+, iberiotoxin, and the
cationic channel blocker Gd3+ but were insensitive to
clofilium, clotrimazole, and apamin. The presence of the recently
cloned swelling-activated, Gd3+-sensitive cation channels
(TRPV4, also known as OTRPC4, TRP12, or VR-OAC) was detected by RT-PCR
in HBE cells. This channel, TRPV4, which senses changes in volume,
might provide the pathway for Ca2+ influx under hypotonic
solutions and, consequently, for the activation of maxi K+ channels.
cell volume regulation; calcium; potassium channels; KCNMA1; airways; 16HBE14o; TRPV4
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INTRODUCTION |
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THE MAINTENANCE of
several epithelial functions (e.g., Cl secretion,
Na+ absorption, and cell volume regulation) requires the
coordinated regulation of apical membrane ion channels [e.g., cystic
fibrosis transmembrane conductance regulator Cl
channel
(CFTR), Ca2+-dependent Cl
channel, and
epithelial Na+ channel (ENaC)] and basolateral membrane
K+ channels (37, 57, 61). A vast account of
the activity and regulation of the epithelial Cl
and
Na+ channels required in such epithelial functions exists
in the literature (4, 5), whereas little is known about
the K+ channels involved in these ion transport processes.
Several types of K+ channels have been identified in
epithelial cells, and their functional roles are just starting to
emerge (14). In the case of the airways, the molecular
identities of the different K+ channels associated with
each type of cellular function remain largely unknown.
In many different types of cells, exposure to hypotonic solutions
induces cell swelling, followed by the return of the cell volume to a
set point close to the original value. This response, known as
regulatory volume decrease (RVD), reflects the cell's adjustments to
the osmotic stress. The response involves the loss of osmolytes and
osmotically obliged water (27, 35). The general mechanism
that allows cells to lose osmolytes has been outlined in many different
cell types, including epithelial cells, and typically employs the
activation of K+ and Cl channels or
transporters, as well as organic osmolyte pathways (27,
35). However, despite the major impetus for the study of
epithelial volume-sensitive Cl
and K+
channels, little advance has been made in the identification of their
molecular nature. Moreover, we do not know whether epithelial cells use
specific volume-sensitive ion channels for cell volume regulatory
purposes or whether they recruit ion channels that are also responsible
for other cellular functions. In the present study we have investigated
the nature of the K+ channels involved in the control of
human airway epithelial volume with special emphasis on their
regulation by increases in intracellular Ca2+ and the
molecular identity of the possible Ca2+ entry pathway.
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METHODS |
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Cells
Human bronchial epithelial cells 16HBE14oSolutions
The isotonic bathing solution contained (mM) 140 NaCl, 2.5 KCl, 1.2 CaCl2, 0.5 MgCl2, 5 glucose, and 10 HEPES, pH 7.25 adjusted with Tris (osmolality: 303 ± 4 mosmol/kgH2O; n = 10 observations). The hypotonic bathing solution (osmolality: 215 ± 8 mosmol/kgH2O; n = 12) was prepared by omitting 50 mM NaCl from the isotonic Hanks' solution and adjusting the osmolality with D-mannitol when necessary. All chemicals were purchased from the Sigma-Aldrich except iberiotoxin (Alomone Laboratories) and clofilium (RBI).Morphometric Analysis
Cell volume experiments were performed at room temperature. Cells were grown on 35-mm cell culture dishes, bathed in isotonic solution, and observed under phase-contrast optics with an inverted microscope (Leica DMIL). The individual cell volume was calculated as described previously (8, 39, 67) and normalized to that measured at time 0 (t = 0).Electrophysiology
Ionic currents were measured by using the whole cell, cell-attached, or excised inside-out recording mode of the patch-clamp technique (24). Strathclyde Electrophysiological Software (written by J. Dempster, University of Strathclyde, Glasgow, Scotland) or pCLAMP8 (Axon Instruments, Foster City, CA) was used for pulse generation, data acquisition through an Axon Digidata analog-to-digital interface, and subsequent analysis. Cells were plated in 35-mm plastic dishes and mounted on the stage of inverted Olympus IX70 or Leica DMIL microscopes.Whole cell ionic currents were measured by using borosilicate glass
electrodes (2-4 M) filled with a solution containing (mM) 140 KCl, 1.2 MgCl2, and 10 HEPES, pH 7.3. The intracellular free Ca2+ concentration
([Ca2+]i), calculated using EqCal (Biosoft,
Cambridge, UK), was adjusted to the desired values by adding different
combinations of CaCl2 and EGTA to the intracellular
solution: 66 nM Ca2+ solution, 0.15 mM CaCl2
and 0.5 mM EGTA; or 200 nM Ca2+ solution, 0.7 mM
CaCl2 and 1 mM EGTA. The osmolality of the intracellular solutions was adjusted to 290 ± 8 mosmol/kgH2O
(n = 23). ATP and GTP were not added to the pipette
solution to delay and reduce the activation of swelling-activated
Cl
channels (7, 19, 22). Isotonic and
hypotonic extracellular solutions were as described above.
Ca2+-free extracellular hypotonic solution (0 [Ca2+]o), containing 0 Ca2+, 1.7 mM MgCl2, and 1 mM EGTA, was also used in several
experiments. Occasionally, the isotonic solution contained 100 mM NaCl
plus 80 mM D-mannitol instead of 140 mM NaCl, and the
hypotonic solution was obtained by omitting D-mannitol.
Whole cell currents were recorded with an Axon 200A amplifier. Cells
were clamped at
80 mV and pulsed for 400 ms from
100 to +100 mV in
20-mV steps. Alternatively, whole cell current-voltage curves were
obtained by applying a ramp of voltage from
120 to +100 mV over a
400-ms period.
Cell-attached single-channel recordings were carried out by using
borosilicate glass electrodes (4-6 M) filled with a solution containing (mM) 100 NaCl, 5 KCl, 1.2 CaCl2, 0.5 MgCl2, 5 glucose, and 10 HEPES, pH 7.25 (osmolality
adjusted with D-mannitol: 300 ± 5 mosmol/kgH2O; n = 15), and the isotonic
bathing solution contained (mM) 105 KCl, 1.2 CaCl2, 0.5 MgCl2, 5 glucose, 10 HEPES, and 80 D-mannitol,
pH 7.25 (osmolality: 305 ± 4 mosmol/kgH2O;
n = 10). The hypotonic bathing solution (osmolality:
215 ± 7 mosmol/kgH2O; n = 10) was
prepared by omitting 80 mM D-mannitol from the isotonic solution. Single-channel currents were obtained by clamping the cells
at different potentials (
20,
40,
60,
80, and
100 mV) for
30 s. Excised inside-out single-channel currents were recorded by
using extracellular (pipette) solutions containing (mM) 140 KCl, 0.7 MgCl2, 10 HEPES, and 5 EDTA, pH 7.24 (osmolality: 302 mosmol/kgH2O). The intracellular (bathing) solution
contained (mM) 140 KCl, 0.7 MgCl2, 10 HEPES, 5 EDTA, and
4.25 CaCl2 (1.6 µM free Ca2+), pH 7.25 (osmolality: 300 mosmol/kgH2O). Currents were low-pass filtered at 1 kHz and sampled at 10 kHz. Single-channel current levels
were measured via all-points histograms, and open probability, Po or NPo, was calculated
from 30-s recordings at the indicated potential in patches containing
only one channel or more than one channel, respectively.
Expression of Maxi K+ and TRPV4
Channels in 16HBE14o Cells
RT-PCR.
Total RNA was extracted from 16HBE14o and HEKhSlo cells [human
embryonic kidney cells permanently transfected with hSlo
(1)] when they were 80% confluent by using the
Nucleospin RNA II kit (Macherey-Nagel), which also degrades the genomic
DNA. Total RNA (1-2 µg) was reverse transcribed to obtain cDNA.
After 2 min of denaturation at 90°C, the RNA was incubated for 1 h at 35°C in 20 µl of reverse transcriptase reaction mix containing
1× RT buffer (Promega), 2 µM oligo(dT12) primer, 125 µM pooled dNTPs, and 200 units of Moloney murine leukemia virus
reverse transcriptase (Promega). One microliter of cDNA product was
sufficient to amplify by PCR a fragment of 479 bp that corresponds to
nucleotides 2675 to 3154 of the hSlo complete coding
sequence (U11717), within a conserved nonalternative spliced region
encompassing the S10 region of the gene, which is not shared with other
known genes (confirmed by a BLASTN search), by using the following
primers: forward 5'-ACCAAGACGATGATGATGACC -3' and reverse
5'-AGCAGAAGATCAGGTCCGTC-3'. The profile was as follows: 95°C for
45 s, 60°C for 1 min, and 72°C for 60 s (30 cycles).
Membrane preparations of 16HBE14o and HEK hSlo cells.
A detailed description of the membrane preparation protocol has been
previously published (23), and only minor modifications have been made to the basic protocol. Briefly, after the cells were
washed with Ca2+- and Mg2+-free PBS, they were
incubated in the same medium with 0.1 mM EGTA at 4°C for 10 min,
scraped, and centrifuged at 800 g for 5 min. The pellet was
then resuspended in (mM) 137 NaCl, 5.6 dextrose, 1 EGTA, and 5 HEPES at
pH 7.4 and centrifuged as above. The pellet was then homogenized in
(mM) 5 Tris · HCl, 5 MgCl2, and 1 EGTA at pH 7.5. The homogenate was centrifuged at 1,000 g
for 5 min, and the supernatant was centrifuged at 100,000 g
for 1 h to obtain crude membranes. Membranes were resuspended in
50 mM Tris · HCl, 5 mM MgCl2, and 1 mM EGTA at pH 7.6 supplemented with 10 µg/ml leupeptin, 10 µg/ml
aprotinin, and 10 µg/ml phenylmethylsulfonyl fluoride and stored at
70°C. Protein content was determined using the Lowry method
(Bio-Rad).
Western blots.
16HBE14o membrane proteins (20 µg) were separated on 8%
SDS-polyacrylamide gels under reducing conditions and
electrotransferred to nitrocellulose paper. Blots were blocked with
TBST (100 mM Tris · HCl, 150 mM NaCl, and 0.1%
Tween 20, at pH 7.5) containing 5% nonfat dry milk overnight at 4°C.
Blots were then incubated with 1:400 affinity-purified anti-maxi K
channel
(913-926)-subunit polyclonal
antibody (62) in TBST-5% nonfat dry milk for 2 h at
room temperature, washed with TBST-5% nonfat dry milk five times for
10 min each time, and then incubated with horseradish peroxidase
(HRP)-conjugated secondary antibody (1:5,000) for 1 h. After being
washed, blots were treated for 10 min with the West Pico
chemiluminescent substrate SuperSignal (Pierce) and autoradiographed on
Amersham Hyperfilm ECL. As a positive control, human embryonic kidney
cells transfected with the maxi K+ channel
-subunit
hSlo (HEK hSlo) displayed a band with a molecular mass of
~120 kDa as expected for the maxi K
-subunit.
[Ca2+]i Measurements
Intracellular Ca2+ concentration measurements were carried out as described previously (56). Cells were incubated in isotonic solution containing 2 µM fura 2-AM (Molecular Probes, Leiden, The Netherlands) for 30 min at room temperature. The cells were then washed thoroughly with isotonic solution. Video microscopic measurements of Ca2+ were obtained by using an Olympus IX70 inverted microscope (Hamburg, Germany) with a ×40 oil-immersion objective (Olympus). The excitation light (340 and 380 nm) was supplied by a Polychrome IV monochromator (Till Photonics, Martinsried, Germany) and directed toward the cells under study by a 505DR dichromatic mirror (Omega Optical, Brattleboro, VT). Fluorescence images were collected by a digital charge-coupled device camera (Hamamatsu Photonics) after being passed through a 535DF emission filter (Omega Optical) by using AquaCosmos software (Hamamatsu Photonics). Fluorescence ratio (340 nm to 380 nm, or 340/380) images were computed every 5 s.Statistics
Results are expressed as means ± SE of n observations. To compare sets of data, we used the Student's t-test provided by SigmaPlot 5 software (Jandel Scientific). Differences were considered statistically significant when P < 0.05. ![]() |
RESULTS |
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Identification of K+ Channels in Human Bronchial Epithelial Cells
Human bronchial epithelial (HBE) cells were routinely clamped at
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The electrophysiological (Fig. 1) and molecular data (Fig. 2) support
the presence of hSlo in HBE cells. To further characterize the nature of the whole cell currents recorded in HBE cells, we used
different inhibitors of K+ channels (Fig.
3), some of which were wide spectrum
[tetraethylammonium (TEA) and Ba2+ (36, 69)]
whereas others were more specific for a particular type of
K+ channel: iberiotoxin and apamin are inhibitory toxins
specific for the large- (maxi K+, or BK) and
small-conductance Ca2+-dependent K+ channels
(SK) (12, 69), respectively; clotrimazole is a well-known inhibitor of intermediate-conductance K+ channels (IK)
(30, 31, 67); and clofilium is an inhibitor of the KCNQ
type of K+ channels (also known as KvLQT) (9).
The latter has been recently shown to be involved in the RVD response
of murine tracheal cells (39). The pharmacological profile
obtained for HBE K+ currents (Fig. 3), i.e., inhibition by
iberiotoxin (n = 6), Ba2+
(n = 6), and TEA (n = 8) but
insensitivity to clofilium (n = 5), apamin
(n = 3), and clotrimazole (n = 5),
could only be explained by the contribution of maxi K+
channels to the generation of such whole cell currents.
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Modulation of Maxi K+ Currents by Hypotonic Cell Swelling
K+ currents were recorded under isotonic conditions in HBE cells dialyzed with solutions containing 66 nM free Ca2+ (Fig. 4A). A substantial increase in K+ currents was achieved within 3 min of exposure of the cell to a 28% hypotonic bathing solution in 19 of 28 cells. The current increase was reversed upon removal of the hypotonic condition (washout). The increase in K+ channel activity following hypotonic shock was inhibited by 100 nM iberiotoxin (Fig. 4B), confirming that the K+ channel activated by hypotonic solutions in HBE cells is a maxi K+ channel. In the presence of iberiotoxin, a small volume-sensitive Cl
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A rise in [Ca2+]i has been implicated in the
cell swelling-mediated modulation of different ion channels (44,
48). To evaluate whether similar mechanisms are operative in the
activation of maxi K+ channels in HBE cells, we recorded
whole cell currents under conditions in which the extracellular
Ca2+ concentration had been modified (Fig.
6). HBE cells dialyzed with a pipette
solution containing 100 nM free Ca2+ and bathed in
isotonic and hypotonic solutions containing 1.2 mM Ca2+
showed an increase in maxi K+ current (Fig. 6A).
Removal of extracellular Ca2+ (0 [Ca2+]o) prevented the increase in maxi
K+ current under hypotonic conditions (Fig. 6B).
Figure 6C shows the time course of
[Ca2+]i in response to hypotonic solutions
measured as the 340/380 fluorescence ratio. The mean increase in the
fluorescence ratio in response to hypotonic solutions was 0.33 ± 0.09 (n = 17). The increase in
[Ca2+]i was mainly due to entry of
extracellular Ca2+, because removal of extracellular
Ca2+ (Fig. 6D) greatly reduced
[Ca2+]i increase (0.07 ± 0.02;
n = 7; P < 0.01). Figure 6E
shows superimposed time courses for the increase in
[Ca2+]i and the activation of maxi
K+ current in response to hypotonic swelling.
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The fact that exposure to hypotonic solutions triggered an increase in
[Ca2+]i (Fig. 6C) and that removal
of extracellular Ca2+ impairs the activation of the maxi
K+ channel (Fig. 6B) suggests the participation
of a Ca2+ entry pathway, activated in response to hypotonic
shocks. This Ca2+ pathway has typically been associated
with stretch-activated ion channels (52, 54). Indeed, the
recent cloning of new members of the family of transient receptor
potential cation channels has provided the molecular basis for at least
one of the pathways for Ca2+ entry following hypotonic
shock (38, 60, 71, 75). To test the involvement of these
cation channels in the activation of RVD mechanisms, we carried out
experiments in the presence of Gd3+, a well-known blocker
of stretch- and swelling-activated cation channels (77).
Figure 7A shows the time
course of a typical experiment in which the holding current and the
current measured at +100 mV were continuously recorded under control
conditions as well as in the presence of Gd3+ and/or a
hypotonic solution. The presence of 100 µM Gd3+ did not
block maxi K+ channels under isotonic conditions (see also
Fig. 7B), but it did prevent the further increase in maxi
K+ channel activity in response to a hypotonic solution.
Upon removal of Gd3+, the maxi K+ current was
readily increased in the presence of a hypotonic solution. Figure
7B shows the mean current-voltage curves obtained from HBE
cells in response to hypotonic shocks in the presence or absence of 100 µM Gd3+. To discard the possibility that the absence of
maxi K+ channel activation could be due to a direct
inhibition of maxi K+ by Gd3+, we measured
single-channel maxi K+ currents using the inside-out
configuration of the patch-clamp technique in the presence or absence
of 100 µM Gd3+ in the pipette and 1.6 µM
Ca2+ in the bathing solution (Fig. 7, C and
D). Neither the channel conductance nor the open probability
was altered in the presence of Gd3+. These results are
consistent with the hypothesis that points to a
Gd3+-sensitive, swelling-activated Ca2+ influx
as the trigger of maxi K+ activation. The recent molecular
identification of a cation channel, TRPV4, which is rapidly activated
under hypotonic conditions when expressed heterologously (38, 60,
71, 75), allowed us to design primers to check its presence in
HBE cells. RT-PCR produced two amplicons of the expected size (see
METHODS), which, after sequencing of the bands, were
confirmed to indeed correspond to the TRPV4 sequence (Fig.
7E).
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RVD in HBE Cells
Relative cell volume changes of HBE cells in response to hypotonic solutions were measured as described in METHODS. Superfusion of HBE cells with a 28% hypotonic solution resulted in a clear increase in cell size, followed by the return to near-original size in isotonic solutions (Fig. 8A). The RVD response observed under control conditions was prevented when the experiments were carried out in the absence of extracellular Ca2+ (Fig. 8B) or in the presence of extracellular Ca2+ and 10 µM Gd3+ (Fig. 8C). To determine whether the activation of maxi K+ channels plays a role in the RVD response of HBE cells, we monitored cell volume changes in response to hypotonic solutions and the effect on RVD of various potassium channel blockers previously tested on the K+ currents of HBE cells (see Fig. 3). Of all the compounds tested, only those shown to block maxi K+ channel activity [5 mM Ba2+ (Fig. 9A), 5 mM TEA (Fig. 9B), and 100 nM iberiotoxin (Fig. 9C)] were effective as inhibitors of the RVD response. On the other hand, 100 nM clotrimazole (n = 3), 1 µM apamin (n = 7), and 100 µM clofilium (n = 4) did not block the RVD response (results not shown).
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DISCUSSION |
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Basolateral K+ channels are critical for epithelial function (14, 16). Recent work carried out in airway epithelia identified K+ channels that are involved in the maintenance of the resting cell membrane potential as well as ion transport processes: intermediate-conductance Ca2+-dependent K+ channels (hIK, KCNN4) (11, 18, 43, 67) and small-conductance K+ channel, known as KvLQT1 (KCNQ1) (6, 70). The latter can form a channel complex with a regulatory subunit, KCNE1 (also named MinK or IsK) or KCNE3 (MiRP2) (6). However, little is known about the regulation and molecular identity of volume-sensitive K+ channels despite their key role in epithelial physiology.
Our results indicate that the 16HBE14o cells possess a K+
current that, on the basis of electrophysiological and pharmacological profiles, is consistent with the activity of Ca2+-dependent
maxi K+ channels (Figs. 1 and 3). No indication of the
presence of functional Ca2+-dependent K+
channels of intermediate conductance [sensitive to clotrimazole (30)], small conductance [sensitive to apamin
(33)], or KvLQT1 channels [sensitive to clofilium
(9)] was obtained in the 16HBE14o
cells. The presence
of maxi K+ channels in 16HBE14o
cells was further
confirmed by RT-PCR and Western blot (Fig. 2).
The present and previous (39, 67) studies have established
that airway epithelial cells undergo an RVD response following a
hypotonic challenge. The RVD response is typically mediated by the loss
of cytosolic K+ and Cl, via the coordinated
activation of K+ and Cl
channels (27,
35). Swelling-activated Cl
channels are present in
most cell types. Their regulation and biophysical properties are well
known, but their molecular identity has been particularly difficult to
resolve, and no clear candidates exist to date (46, 59, 63,
74).
Different swelling-sensitive K+ channels have been described in several cell preparations (48, 58, 72), although according to Hoffmann and Dunham (27), "there are no K+ channels which are directly activated by swelling, meaning that activation of K+ channels following cell swelling is secondary to the membrane depolarization or the production of intracellular signals." Exceptions to that statement are the K+ channels directly activated by membrane stretch (2, 53).
Among the numerous K+ channels associated with the mechanisms of cell volume control, only a handful have been identified at the molecular level: 1) voltage-gated K+ channels, including Kv1.3 (17), Kv1.5 (21), and KCNQ/KCNE complex (10, 39); 2) background TREK-1 (49), TRAAK (41), and TASK-2 (45) channels, with TREK-1 and TRAAK being mechanosensitive; and 3) Ca2+-dependent K+ channels of large (2) and intermediate conductance (67).
Cell volume regulation following hypotonic or isotonic swelling in epithelial cells is normally associated with changes in intracellular Ca2+ concentrations (44, 48). However, the source of Ca2+ appears to be different, depending on the original stimulus, i.e., hypotonic cell swelling typically involves extracellular Ca2+, whereas isotonic swelling following nutrient absorption involves mobilization of intracellular Ca2+ (40), although descriptions of Ca2+ release from intracellular stores under hypotonic conditions also exist (29, 76). Our results with HBE cells showing no increase of intracellular Ca2+ in Ca2+-free hypotonic conditions resembled those reported by Ishii et al. (29) on Intestine 407 epithelial cells. On the contrary, RVD in nonepithelial cells normally shows no Ca2+ dependence (3).
As we discussed earlier, the RVD mechanisms in most cells involve the
coordinated activation of Cl and K+ channels.
The swelling-activated Cl
channels are characteristically
Ca2+ independent. Exceptions include swelling-sensitive
Cl
channels, the activation of which depends on
extracellular or intracellular Ca2+ concentration
(34, 51, 68) or is prevented by high
[Ca2+]i (25). Therefore, it is
plausible to think that in those epithelial cells showing a
Ca2+-dependent RVD, the Ca2+-dependency is
typically conferred by the K+ channels (see Ref.
48 for a recent review). Ca2+- and
swelling-modulated K+ channels have been characterized in
epithelia that also show Ca2+-dependent RVD (26, 50,
55, 67). Among the Ca2+-dependent K+
channels responding to cell swelling, maxi K+ channels have
been identified in several epithelia, including proximal tubule of the
kidney (20, 32), lacrimal gland (47), and
airways (present study) as well as nonepithelial cells
(73).
The activation of maxi K+ channels following hypotonic
swelling of the 16HBE14o cells is accompanied by increases in
[Ca2+]i, requires extracellular
Ca2+ (Fig. 6), and is blocked by Gd3+, a
blocker of stretch- and/or swelling-activated cation channels (Fig. 7).
Similarly, RVD in 16HBE14o
cells is inhibited in the absence of
extracellular Ca2+ or in the presence of Gd3+
(Fig. 8). These observations suggest that the mechanisms activated to
achieve a complete RVD consist of an increase in Ca2+
entry, most likely via stretch- or swelling-activated cation channels,
the nature of which has only started to emerge (see below), and the
subsequent activation of maxi K+ channels, a mechanism
previously characterized for other cell preparations (13,
28).
Several laboratories have identified a new TRP channel (TRPV4) that is responsive to changes in extracellular osmolarity (38, 60, 71, 75). RT-PCR experiments confirmed the presence of TRPV4 amplicons in the HBE cells (Fig. 7E). Therefore, this TRPV4 channel could provide the Ca2+ influx pathway associated with maxi K+ channel activation and RVD response in HBE cells.
A similar series of events appears to underlie the activation of the
RVD response in the human tracheal CFT1-LCFSN cells (67), although in this case the Ca2+-dependent K+
channel activated is of intermediate conductance (hIK, also known as
KCNN4). The fact that 16HBE14o cells (present study) and CFT1-LCFSN cells use different Ca2+-dependent K+ channels
for the RVD response might be related to their different origin, with
the former being bronchial and the later tracheal cells. It would be
interesting to show whether native human airway tissue shows similar
regional differences. The only study addressing the RVD capability of
native airway epithelial cells was carried out on murine tracheal cells
(39). That study demonstrated that in murine tracheal
cells, the pathway for K+ efflux was via a
clofilium-sensitive KCNQ K+ channel.
In summary, the human bronchial epithelial cell line 16HBE14o shows
Ca2+-dependent RVD. The swelling of 16HBE14o
triggers
extracellular Ca2+ entry, most likely through stretch- or
swelling-activated cation channels, with TRPV4 being a strong candidate
for the Ca2+ pathway, and the activation of
Ca2+-dependent maxi K+ channels. The
identification of the mechanisms involved in cell volume regulation in
epithelial cells is relevant to the pathophysiology of cystic fibrosis
because it has been shown that RVD is impaired in both cystic fibrosis
murine intestine (65, 66) and human airways
(67).
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ACKNOWLEDGEMENTS |
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We thank Dr. D. Gruenert (University of Vermont, Burlington, VT) for the gift of HBE cells and Dr. L. Toro for the gift of anti-Maxi K+ antibody. We also acknowledge the help of S. Thambapillai with the initial cell volume experiments, M. I. Bahamonde with the membrane preparations, and H. Lock for comments on the manuscript.
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FOOTNOTES |
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* J. M. Fernandez-Fernandez and M. Nobles contributed equally to this work.
This work was supported by the Spanish Ministry of Science and Technology and the Human Frontiers Science Program.
Address for reprint requests and other correspondence: M. A. Valverde, Departament de Ciències Experimentals i de la Salut, Universitat Pompeu Fabra, C/ Dr. Aiguader 80, 08003 Barcelona, Spain (E-mail: miguel.valverde{at}cexs.upf.es).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
August 14, 2002;10.1152/ajpcell.00245.2002
Received 28 May 2002; accepted in final form 31 July 2002.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Ahring, PK,
Strobaek D,
Christophersen P,
Olesen SP,
and
Johansen TE.
Stable expression of the human large-conductance Ca2+-activated K+ channel alpha-and beta-subunits in HEK293 cells.
FEBS Lett
415:
67-70,
1997[ISI][Medline].
2.
Allard, B,
Couble ML,
Magloire H,
and
Bleicher F.
Characterization and gene expression of high conductance calcium-activated potassium channels displaying mechanosensitivity in human odontoblasts.
J Biol Chem
275:
25556-25561,
2000
3.
Altamirano, J,
Brodwick MS,
and
Alvarez-Leefmans FJ.
Regulatory volume decrease and intracellular Ca2+ in murine neuroblastoma cells studied with fluorescent probes.
J Gen Physiol
112:
145-160,
1998
4.
Alvarez de la Rosa, D,
Canessa CM,
Fyfe GK,
and
Zhang P.
Structure and regulation of amiloride-sensitive sodium channels.
Annu Rev Physiol
62:
573-594,
2000[ISI][Medline].
5.
Anderson, MP,
Sheppard DN,
Berger HA,
and
Welsh MJ.
Chloride channels in the apical membrane of normal and cystic fibrosis airway and intestinal epithelia.
Am J Physiol Lung Cell Mol Physiol
263:
L1-L14,
1992
6.
Bleich, M,
and
Warth R.
The very small-conductance K+ channel KvLQT1 and epithelial function.
Pflügers Arch
440:
202-206,
2000[ISI][Medline].
7.
Bond, T,
Basavappa S,
Christensen M,
and
Strange K.
ATP dependence of the ICl,swell channel varies with rate of cell swelling. Evidence for two modes of channel activation.
J Gen Physiol
113:
441-456,
1999
8.
Bond, TD,
Ambikapathy S,
Mohammad S,
and
Valverde MA.
Osmosensitive Cl currents and their relevance to regulatory volume decrease in human intestinal T-84 cells: outwardly vs. inwardly rectifying currents.
J Physiol
511:
45-54,
1998
9.
Busch, AE,
and
Suessbrich H.
Role of the ISK protein in the IminK channel complex.
Trends Pharmacol Sci
18:
26-29,
1997[ISI][Medline].
10.
Busch, AE,
Varnum M,
Adelman JP,
and
North RA.
Hypotonic solution increases the slowly activating potassium current IsK expressed in Xenopus oocytes.
Biochem Biophys Res Commun
184:
804-810,
1992[ISI][Medline].
11.
Butt, AG,
Clapp WL,
and
Frizzell RA.
Potassium conductances in tracheal epithelium activated by secretion and cell swelling.
Am J Physiol Cell Physiol
258:
C630-C638,
1990
12.
Castle, NA,
Haylett DG,
and
Jenkinson DH.
Toxins in the characterization of potassium channels.
Trends Neurosci
12:
59-65,
1989[ISI][Medline].
13.
Christenson, O.
Mediation of cell volume regulation by Ca2+ influx through stretch-activated channels.
Nature
330:
66-68,
1987[ISI][Medline].
14.
Cotton, CU.
Basolateral potassium channels and epithelial ion transport.
Am J Respir Cell Mol Biol
23:
270-272,
2000
15.
Cozens, AL,
Yezzi MJ,
Kunzelmann K,
Ohrui T,
Chin L,
Eng K,
Finkbeiner WE,
Widdicombe JH,
and
Gruenert DC.
CFTR expression and chloride secretion in polarized immortal human bronchial epithelial cells.
Am J Respir Cell Mol Biol
10:
38-47,
1994[Abstract].
16.
Dawson, DC,
and
Richards NW.
Basolateral K+ conductance: role in regulation of NaCl absorption and secretion.
Am J Physiol Cell Physiol
259:
C181-C195,
1990
17.
Deutsch, C,
and
Chen LQ.
Heterologous expression of specific K+ channels in T lymphocytes: functional consequences for volume regulation.
Proc Natl Acad Sci USA
90:
10036-10040,
1993[Abstract].
18.
Devor, DC,
Singh AK,
Lambert LC,
DeLuca A,
Frizzell RA,
and
Bridges RJ.
Bicarbonate and chloride secretion in Calu-3 human airway epithelial cells.
J Gen Physiol
113:
743-760,
1999
19.
Diaz, M,
Valverde MA,
Higgins CF,
Rucareaunu C,
and
Sepulveda FV.
Volume-activated chloride channels in HeLa cells are blocked by verapamil and dideoxyforskolin.
Pflügers Arch
422:
347-353,
1993[ISI][Medline].
20.
Dube, L,
Parent L,
and
Sauve R.
Hypotonic shock activates a maxi K+ channel in primary cultured proximal tubule cells.
Am J Physiol Renal Fluid Electrolyte Physiol
259:
F348-F356,
1990
21.
Felipe, A,
Snyders DJ,
Deal KK,
and
Tamkun MM.
Influence of cloned voltage-gated K+ channel expression on alanine transport, Rb+ uptake, and cell volume.
Am J Physiol Cell Physiol
265:
C1230-C1238,
1993
22.
Gill, DR,
Hyde SC,
Higgins CF,
Valverde MA,
Mintenig GM,
and
Sepulveda FV.
Separation of drug transport and chloride channel functions of the human multidrug resistance P-glycoprotein.
Cell
71:
23-32,
1992[ISI][Medline].
23.
Graeser, D,
and
Neubig RR.
Methods for the study of receptor/G-protein interaction.
In: Signal Transduction: A Practical Approach, edited by Milligan G.. Oxford, UK: IRL, 1992, p. 1-29.
24.
Hamill, OP,
Marty A,
Neher E,
Sakmann B,
and
Sigworth J.
Improved patch-clamp techniques for high resolution current recording from cells and cell-free membrane patches.
Pflügers Arch
391:
85-100,
1981[ISI][Medline].
25.
Hardy, SP,
Goodfellow HR,
Valverde MA,
Gill DR,
Sepulveda FV,
and
Higgins CF.
Protein kinase C-mediated phosphorylation of the human multidrug-resistance P-glycoprotein regulates cell volume-activated chloride channels.
EMBO J
14:
68-75,
1995[Abstract].
26.
Hazama, A,
and
Okada Y.
Ca2+ sensitivity of volume-regulatory K+ and Cl channels in cultured human epithelial cells.
J Physiol
402:
687-702,
1988[Abstract].
27.
Hoffmann, EK,
and
Dunham PB.
Membrane mechanisms and intracellular signalling in cell volume regulation.
Int Rev Cytol
161:
173-262,
1995[ISI][Medline].
28.
Hoyer, J,
Distler A,
Haase W,
and
Gogelein H.
Ca2+ influx through stretch-activated cation channels activates maxi K+ channels in porcine endocardial endothelium.
Proc Natl Acad Sci USA
91:
2367-2371,
1994[Abstract].
29.
Ishii, T,
Hashimoto T,
and
Ohmori H.
Hypotonic stimulation induced Ca2+ release from IP3-sensitive internal stores in a green monkey kidney cell line.
J Physiol
493:
371-384,
1996[Abstract].
30.
Ishii, TM,
Silvia C,
Hirschberg B,
Bond CT,
Adelman JP,
and
Maylie J.
A human intermediate conductance calcium-activated potassium channel.
Proc Natl Acad Sci USA
94:
11651-11656,
1997
31.
Joiner, WJ,
Wang LY,
Tang MD,
and
Kaczmarek LK.
hSK4, a member of a novel subfamily of calcium-activated potassium channels.
Proc Natl Acad Sci USA
94:
11013-11018,
1997
32.
Kawahara, K,
Ogawa A,
and
Suzuki M.
Hyposmotic activation of Ca-activated K channels in cultured rabbit kidney proximal tubule cells.
Am J Physiol Renal Fluid Electrolyte Physiol
260:
F27-F33,
1991
33.
Kohler, M,
Hirschberg B,
Bond CT,
Kinzie JM,
Marrion NV,
Maylie J,
and
Adelman JP.
Small-conductance, calcium-activated potassium channels from mammalian brain.
Science
273:
1709-1714,
1996
34.
Kotera, T,
and
Brown PD.
Calcium-dependent chloride current activated by hyposmotic stress in rat lacrimal acinar cells.
J Membr Biol
134:
67-74,
1993[ISI][Medline].
35.
Lang, F,
Busch GL,
Ritter M,
Volkl H,
Waldegger S,
Gulbins E,
and
Haussinger D.
Functional significance of cell volume regulatory mechanisms.
Physiol Rev
78:
247-306,
1998
36.
Latorre, R,
Oberhauser A,
Labarca P,
and
Alvarez O.
Varieties of Ca2+-activated K+ channels.
Annu Rev Physiol
51:
385-399,
1989[ISI][Medline].
37.
Liedtke, CM.
Regulation of chloride transport in epithelia.
Annu Rev Physiol
51:
143-160,
1989[ISI][Medline].
38.
Liedtke, W,
Choe Y,
Marti-Renom MA,
Bell AM,
Denis CS,
Sali A,
Hudspeth AJ,
Friedman JM,
and
Heller S.
Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor.
Cell
103:
525-535,
2000[ISI][Medline].
39.
Lock, H,
and
Valverde MA.
Contribution of the IsK (MinK) potassium channel subunit to regulatory volume decrease in murine tracheal epithelial cells.
J Biol Chem
275:
34849-34852,
2000
40.
MacLeod, RJ.
How an epithelial cell swells is determinant of the signaling pathways that activate RVD.
In: Cellular and Molecular Physiology of Cell Volume Regulation, edited by Strange K.. Boca Raton, FL: CRC, 1994, p. 191-213.
41.
Maingret, F,
Fosset M,
Lesage F,
Lazdunski M,
and
Honore E.
TRAAK is a mammalian neuronal mechano-gated K+ channel.
J Biol Chem
274:
1381-1387,
1999
42.
Marijic, J,
Li Q,
Song M,
Nishimaru K,
Stefani E,
and
Toro L.
Decreased expression of voltage- and Ca2+-activated K+ channels in coronary smooth muscle during aging.
Circ Res
88:
210-216,
2001
43.
McCann, JD,
Matsuda J,
Garcia M,
Kaczorowski G,
and
Welsh MJ.
Basolateral K+ channels in airway epithelia. I. Regulation by Ca2+ and block by charybdotoxin.
Am J Physiol Lung Cell Mol Physiol
258:
L334-L342,
1990
44.
McCarty, NA,
and
O'Neil RG.
Calcium signalling in cell volume regulation.
Physiol Rev
72:
1037-1061,
1992
45.
Niemeyer, MI,
Cid LP,
Barros LF,
and
Sepulveda FV.
Modulation of the two-pore domain acid-sensitive K+ channel TASK-2 (KCNK5) by changes in cell volume.
J Biol Chem
276:
43166-43174,
2001
46.
Okada, Y.
Volume expansion-sensing outward-rectifier Cl channel: fresh start to the molecular identity and volume sensor.
Am J Physiol Cell Physiol
273:
C755-C789,
1997
47.
Park, KP,
Beck JS,
Douglas IJ,
and
Brown PD.
Ca2+-activated K+ channels are involved in regulatory volume decrease in acinar cells isolated from the rat lacrimal gland.
J Membr Biol
141:
193-201,
1994[ISI][Medline].
48.
Pasantes-Morales, H,
and
Morales-Mulia S.
Influence of calcium on regulatory volume decrease: role of potassium channels.
Nephron
86:
414-427,
2000[ISI][Medline].
49.
Patel, AJ,
Honore E,
Maingret F,
Lesage F,
Fink M,
Duprat F,
and
Lazdunski M.
A mammalian two pore domain mechano-gated S-like K+ channel.
EMBO J
17:
4283-4290,
1998
50.
Roman, RM,
Wang Y,
and
Fitz JG.
Regulation of cell volume in human biliary cell line: activation of K+ and Cl currents.
Am J Physiol Gastrointest Liver Physiol
271:
G239-G248,
1996
51.
Rubera, I,
Tauc M,
Poujeol C,
Bohn MT,
Bidet M,
De Renzis G,
and
Poujeol P.
Cl and K+ conductances activated by cell swelling in primary cultures of rabbit distal bright convoluted tubules.
Am J Physiol Renal Physiol
273:
F680-F697,
1997
52.
Sachs, F,
and
Morris CE.
Mechanosensitive ion channels in nonspecialized cells.
Rev Physiol Biochem Pharmacol
132:
1-77,
1998[ISI][Medline].
53.
Sackin, H.
A stretch-activated K+ channel sensitive to cell volume.
Proc Natl Acad Sci USA
86:
1731-1753,
1989[Abstract].
54.
Sackin, H.
Stretch-activated ion channels.
Kidney Int
48:
1134-1147,
1995[ISI][Medline].
55.
Samman, G,
Ohtsuyama M,
Sato F,
and
Sato K.
Volume-activated K+ and Cl pathways of dissociated eccrine clear cells.
Am J Physiol Regul Integr Comp Physiol
265:
R990-R1000,
1993
56.
Scheenen, WJ,
Hofer AM,
and
Pozzan T.
Intracellular measurement of calcium using fluorescent probes.
In: Cell Biology. A Laboratory Handbook, edited by Celis JE.. San Diego, CA: Academic, 1998, p. 363-374.
57.
Schultz, SG.
Homocellular regulatory mechanisms in sodium-transporting epithelia: avoidance of extinction by "flush-through".
Am J Physiol Renal Fluid Electrolyte Physiol
241:
F579-F590,
1981
58.
Schultz, SG,
Dubinsky WP,
and
Lapointe JY.
Volume regulation and "cross-talk" in sodium-absorbing epithelial cells.
Contrib Nephrol
123:
205-219,
1998[ISI][Medline].
59.
Strange, K,
Emma F,
and
Jackson PS.
Cellular and molecular physiology of volume-sensitive anion channels.
Am J Physiol Cell Physiol
270:
C711-C730,
1996
60.
Strotmann, R,
Harteneck C,
Nunnenmacher K,
Schultz G,
and
Plant TD.
OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity.
Nat Cell Biol
2:
695-702,
2000[ISI][Medline].
61.
Sullivan, SK,
and
Field M.
Ion transport across mammalian small intestine.
In: Handbook of Physiology. The Gastrointestinal System. Absorption and Secretion. Bethesda, MD: Am. Physiol. Soc, 1991, sect. 6, vol. IV, chapt. 10, p. 287-301.
62.
Tanaka, Y,
Meera P,
Song M,
Knaus HG,
and
Toro L.
Molecular constituents of maxi KCa channels in human coronary smooth muscle: predominant alpha + beta subunit complexes.
J Physiol
502:
545-557,
1997[Abstract].
63.
Valverde, MA.
ClC channels: leaving the dark ages on the verge of a new millennium.
Curr Opin Cell Biol
11:
509-516,
1999[ISI][Medline].
64.
Valverde, MA,
Hardy SP,
and
Sepulveda FV.
Chloride channels: a state of flux.
FASEB J
9:
509-515,
1995
65.
Valverde, MA,
O'Brien JA,
Sepulveda FV,
Ratcliff RA,
Evans MJ,
and
College WH.
Impaired cell volume regulation in intestinal crypt epithelia of cystic fibrosis mice.
Proc Natl Acad Sci USA
92:
9038-9041,
1995[Abstract].
66.
Valverde, MA,
Vázquez E,
Munoz FJ,
Nobles M,
Delaney SJ,
Wainwright BJ,
Colledge WH,
and
Sheppard DN.
Murine CFTR channel and its role in regulatory volume decrease of small intestine crypts.
Cell Physiol Biochem
10:
321-328,
2000[ISI][Medline].
67.
Vázquez, E,
Nobles M,
and
Valverde MA.
Defective regulatory volume decrease in human cystic fibrosis tracheal cells because of altered regulation of intermediate conductance Ca2+-dependent potassium channels.
Proc Natl Acad Sci USA
98:
5329-5334,
2001
68.
Verdon, B,
Winpenny JP,
Whitfield KJ,
Argent BE,
and
Gray MA.
Volume-activated chloride currents in pancreatic duct cells.
J Membr Biol
147:
173-183,
1995[ISI][Medline].
69.
Vergara, C,
Latorre R,
Marrion NV,
and
Adelman JP.
Calcium-dependent potassium channels.
Curr Opin Neurobiol
8:
321-329,
1998[ISI][Medline].
70.
Wang, Q,
Curran ME,
Splawski I,
Burn TC,
Millholland JM,
VanRaay TJ,
Shen J,
Timothy KW,
Vincent GM,
de Jager T,
Schwartz PJ,
Toubin JA,
Moss AJ,
Atkinson DL,
Landes GM,
Connors TD,
and
Keating MT.
Positional cloning of a novel potassium channel gene: KVLQT1 mutations cause cardiac arrhythmias.
Nat Genet
12:
17-23,
1996[ISI][Medline].
71.
Watanabe, H,
Davis JB,
Smart D,
Jerman JC,
Smith GD,
Hayes P,
Vriens J,
Cairns W,
Wissenbach U,
Prenen J,
Flockerzi V,
Droogmans G,
Benham CD,
and
Nilius B.
Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives.
J Biol Chem
277:
13569-13577,
2002
72.
Wehner, F.
Cell volume-regulated cation channels.
Contrib Nephrol
123:
8-20,
1998[ISI][Medline].
73.
Weskamp, M,
Seidl W,
and
Grissmer S.
Characterization of the increase in [Ca2+]i during hypotonic shock and the involvement of Ca2+-activated K+ channels in the regulatory volume decrease in human osteoblast-like cells.
J Membr Biol
178:
11-20,
2000[ISI][Medline].
74.
Weylandt, KH,
Valverde MA,
Nobles M,
Raguz S,
Amey JS,
Diaz M,
Nastrucci C,
Higgins CF,
and
Sardini A.
Human ClC-3 is not the swelling-activated chloride channel involved in cell volume regulation.
J Biol Chem
276:
17461-17467,
2001
75.
Wissenbach, U,
Bodding M,
Freichel M,
and
Flockerzi V.
Trp12, a novel Trp related protein from kidney.
FEBS Lett
485:
127-134,
2000[ISI][Medline].
76.
Wu, X,
Yang H,
Iserovich P,
Fischbarg J,
and
Reinach PS.
Regulatory volume decrease by SV40-transformed rabbit corneal epithelial cells requires ryanodine-sensitive Ca2+-induced Ca2+ release.
J Membr Biol
158:
127-136,
1997[ISI][Medline].
77.
Yang, XC,
and
Sachs F.
Block of stretch-activated ion channels in Xenopus oocytes by gadolinium and calcium ions.
Science
243:
1068-1071,
1989[ISI][Medline].