AJP - Lung AJP: Renal Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Lung Cell Mol Physiol 292: L1432-L1443, 2007. First published February 23, 2007; doi:10.1152/ajplung.00032.2007
1040-0605/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
292/6/L1432    most recent
00032.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (2)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Fischer, H.
Right arrow Articles by Widdicombe, J. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Fischer, H.
Right arrow Articles by Widdicombe, J. H.

Basolateral Cl channels in primary airway epithelial cultures

Horst Fischer,1 Beate Illek,1 Walter E. Finkbeiner,2 and Jonathan H. Widdicombe3

1Children's Hospital Oakland Research Institute, Oakland, 2Department of Pathology, University of California, San Francisco, and 3Department of Physiology and Membrane Biology, University of California, Davis, California

Submitted 22 January 2007 ; accepted in final form 21 February 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Salt and water absorption and secretion across the airway epithelium are important for maintaining the thin film of liquid lining the surface of the airway epithelium. Movement of Cl across the apical membrane involves the CFTR Cl channel; however, conductive pathways for Cl movement across the basolateral membrane have been little studied. Here, we determined the regulation and single-channel properties of the Cl conductance (GCl) in airway surface epithelia using epithelial cultures from human or bovine trachea and freshly isolated ciliated cells from the human nasal epithelium. In Ussing chamber studies, a swelling-activated basolateral GCl was found, which was further stimulated by forskolin and blocked by N-phenylanthranilic acid (DPC) = sucrose > flufenamic acid = niflumic acid = glibenclamide > CdCl2 = 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) = DIDS = ZnCl2 > tamoxifen > 4,4'-dinitro-2,2'-stilbene-disulfonate disodium salt (DNDS). In whole cell patch-clamp experiments, three types of GCl were identified: 1) a voltage-activated, DIDS- (but not Cd-) blockable and osmosensitive GCl; 2) an inwardly rectifying, hyperpolarization-activated and Cd-sensitive GCl; and 3) a forskolin-activated, linear GCl, which was insensitive to Cd and DIDS. In cell-attached patch-clamp recordings, the basolateral pole of isolated ciliated cells expressed three types of Cl channels: 1) an outwardly rectifying, swelling-activated Cl channel; 2) a strongly inwardly rectifying Cl channel; and 3) a forskolin-activated, low-conductance channel. We propose that, depending on the driving force for Cl across the apical membrane, basolateral Cl channels confine Cl– secretion or support transcellular Cl– absorption.

airway epithelium; ion transport; absorption; secretion; chloride channels; basolateral membrane

SALT AND WATER TRANSPORT BY the airway epithelium is critically important in determining the volume of the airway surface liquid. Cl may be secreted or absorbed across the epithelium depending on its electrochemical driving force across the apical membrane, which in turn is determined by other ion transport processes, most notably active Na absorption (19, 46). Basolateral Cl channels have been investigated in other epithelia (4, 13, 14, 21, 22, 43) where roles in Cl absorption and secretion and in cell volume regulation have been suggested. Studies in the airway epithelium have largely focused on the apical membrane, which is dominated by the CFTR-mediated Cl conductance (47). However, early studies suggested a basolateral Cl conductance (GCl) in airway epithelium (40, 41), which was further investigated more recently (15, 32, 35). In principle, a basolateral GCl is expected to support transcellular Cl absorption or limit Cl secretion.

The CFTR Cl channel has been shown to be localized predominantly in apical membranes. Several other GCl and Cl channel types have been described in whole cell patch-clamp recordings. Outwardly rectifying whole cell Cl conductances that are regulated by intracellular Ca and volume have been found in cultured human nasal cells (11). Using ciliated mouse nasal epithelial cells, Tarran et al. (33) have identified three different types of whole cell Cl conductances: a hyperpolarization-activated GCl, a voltage-activating GCl, and a minor voltage-independent GCl. However, whether they are localized in the apical or basolateral plasma membrane is unclear.

In the present work, we assessed the localization and biophysical properties of Cl channels in the basolateral membrane of airway surface epithelial cells. In contrast to previous work, we combined transepithelial recordings from apically nystatin-permeabilized tracheal epithelia with both whole cell and single-channel patch-clamp recordings. We identified three biophysically distinct basolateral GCl in airway epithelia. We propose that the basolateral GCl is a determinant of the direction and size of Cl movement across the airway epithelium.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. Human and bovine tracheal primary cultures were isolated and cultured as previously described (20, 46). In brief, strips of epithelium were removed from the underlying tissues and treated with protease overnight. Cells were plated (~106 cells/cm2) on permeable filter supports (0.4-µm pore size, 1-cm2 area, Transwell; Corning Costar, Cambridge, MA) precoated with human placental collagen (15 µg/cm2). Human tracheal cells were grown in a 1:1 mixture of Dulbecco's modified Eagle's medium and Ham's F-12 nutrient medium (DMEM/F-12) supplemented with 2% Ultroser G (Pall). Bovine tracheal cells were grown in DMEM/F-12 culture medium containing the following growth factors: 10 µg/ml insulin, 5 µg/ml transferrin, 20 ng/ml triiodothyronine, 0.36 µg/ml hydrocortisone, 7.5 mg/ml endothelial cell growth supplement, 25 ng/ml epidermal growth factor, and 10–7 M retinoic acid. All media used in cell isolation and primary culture contained the following antibiotics: 100 U/ml penicillin, 100 µg/ml streptomycin, 2.5 mg/ml fungizone, and 100 µg/ml gentamicin. Tracheal sheets were grown to confluence in an air-liquid interface in a tissue culture incubator gassed with 5% CO2 and air. Transepithelial potential (Vte) and transepithelial resistance (Rte) were monitored routinely under sterile condition using an EVOM voltmeter (World Precision Instruments). Cell sheets were used for transepithelial experiments when Vte was larger than –30 mV.

Transepithelial measurements. Tracheal sheets grown on filters were mounted in Ussing chambers; Vte was clamped to zero and pulsed every 20 or 50 s to 2 mV to monitor the Rte as described in detail in Ref. 18. Positive values of short-circuit current (Isc) represent net transfer of negative charge into the lumen. Initially, tissues were bathed in symmetrical Ringer solution, and Isc was allowed to stabilize. Then, a mucosal-to-serosal Cl gradient was applied, and the apical membrane was permeabilized with nystatin (100–300 µM). The mucosal high Cl solution contained, in mM, 120 NaCl, 20 NaHCO3, 5 KHCO3, 1.3 NaH2PO4, 1.5 CaCl2, 1.0 MgCl2, pH 7.4. In the serosal low Cl solutions, NaCl was exchanged for Na gluconate. In HEPES-buffered solutions, all HCO3 salts were replaced by respective Cl salts, and 10 mM HEPES was added. In all experiments, lack of change in Isc on addition of amiloride (10 µM, mucosal) was taken as evidence of successful permeabilization. Currents generated by the Na-K-ATPase were blocked with ouabain (50 µM, serosal).

Blocker kinetics of fast-acting Cl channel blockers were quantified by addition of stepwise increasing concentrations. Steady-state currents for each drug concentration were measured and fitted to a Michaelis-Menten function of the form Isc = (Imax x CnH)/(KBnH + CnH) by nonlinear regression, where Imax is the maximally blocked current, C is the drug concentration, KB is the half-maximal blocker concentrations, and nH is the Hill coefficient.

Patch-clamp experiments. The patch-clamp procedure has been described in detail (8, 16). Experiments were done on freshly isolated human nasal epithelial cells obtained by a brushing procedure approved by the Internal Review Board of Children's Hospital Oakland. A sterile cytology brush (3-mm diameter, 12-mm length; Mill-Rose Laboratories, Mentor, OH) was inserted ~2 cm into one nostril of a healthy volunteer, and the floor and lateral wall of the inferior turbinate was brushed for ~10 s. The brush was washed with 1 ml of cell culture medium. Single nasal epithelial cells were identified under a microscope by their beating cilia and used for patch-clamp experiments within 2 h of isolation. Cells were immobilized by placing them on the bottom of a patch-clamp chamber coated with poly-L-lysine. Patch pipettes were sealed on the basal half of the basolateral pole, which was readily identifiable.

Whole cell recordings were done with symmetrical N-methyl-D-glucamine chloride (NMDG-Cl) solutions, and the membrane potential (Vm) was clamped to a negative holding potential. In some experiments, voltage pulses were overlaid on Vm to monitor the membrane conductance. Bath solution contained, in mM, 145 NMDG-Cl, 1.7 CaCl2, 1 MgCl2, 20 HEPES, 20 glucose, pH 7.3 adjusted with NMDG. In the whole cell mode, cell swelling was provoked by reducing the concentrations of HEPES and glucose to 2 mM. The pipette solution contained, in mM, 148 NMDG-Cl, 5 EGTA, 1 MgCl2, 0.5 HEPES, 0.5 glucose, 5 MgATP, 0.1 NaGTP, pH 7.3 adjusted with NMDG. Current-voltage (I-V) step protocols were used to identify voltage-dependent characteristics of the investigated conductances. I-V relation shown in the figures represent difference currents calculated by subtracting currents recorded at two consecutive time points (e.g., before and after addition of blocker). Steady-state I-V relations were constructed by averaging 10 samples at the end of each pulse. Sample frequency was 2 kHz. Chord conductances at –100 mV and 100 mV were used to quantify GCl of rectifying I-V relationships. Membrane capacitance (Cm) was estimated from the current transient induced by a 20-mV voltage pulse.

Single-channel recordings were done in the cell-attached mode on the basolateral pole of ciliated cells. The reported holding potentials are referenced to the pipette. In the cell-attached mode, pipette and bath solutions contained, in mM, 145 NMDG-Cl, 1.7 CaCl2, 1 MgCl2, 20 HEPES, 20 glucose, pH 7.3 adjusted with NMDG. Single-channel conductances (g) of rectifying channels are reported as the chord conductances at +100 mV and –100 mV. The voltage dependence of the single-channel current (i) and the open probability (Po) were calculated from recordings where the holding potential was clamped in 25-mV and 2-s steps from –100 mV to +100 mV. Voltage step protocols were applied 2–4 times in succession, and values determined for i and Po were averaged. The time-averaged single-channel current was determined from the product of i x Po for every recording. Single-channel recordings were sampled at 2 kHz and filtered at 500 Hz. For display in some figures, the sampling and filter frequencies were further reduced as noted. All patch-clamp recordings were done at 35–37°C.

Chemicals. The following drugs were used: flufenamic acid (FFA; 1 M in DMSO, used at 1 mM); niflumic acid (NFA; 1 M in DMSO, used at 3 mM); glibenclamide (300 mM in DMSO after sonication, used at 500 µM); DIDS (100 mM in DMSO, used at 100 µM); 4,4'-dinitro-2,2'-stilbene-disulfonate disodium salt (DNDS; 500 mM in DMSO, used at 500 µM); N-phenylanthranilic acid (DPC; 200 mM in ethanol, used at 5 mM; Aldrich); forskolin (20 mM in DMSO, used at 20 µM; Calbiochem); 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB; 100 mM in DMSO, used at 100 µM; Research Biochem); tamoxifen (50 mM in DMSO, used at 50 µM); CdCl2 and ZnCl2 (100 mM in water, used at 100 µM); and nystatin (100 mM in DMSO, used at 100 µM to 300 µM). Sucrose was dissolved at final concentrations directly in solutions.

Statistics. Data are reported as original or as means ± SE. Average values were compared using factorial ANOVA followed by t-tests. One-sample t-tests were used to determine significant differences from zero. If normality tests failed, nonparametric signed-rank tests were used instead. Calculated P values are given, and P < 0.05 was considered significant. All statistical calculations were done with StatView v4.5 (Abacus Concepts) or SigmaStat v3.5 (SYSTAT Software).


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Identification of the basolateral Cl conductance in human and bovine primary tracheal epithelia. Most experimental protocols were performed on both human and bovine cultures. However, owing to the limited supply of human cultures, some experimental sets were performed only in bovine cultures, as noted.

To study the basolateral membrane in Ussing chambers, a 125-to-5 mM mucosal-to-serosal Cl gradient was applied, and the apical membrane was permeabilized with nystatin (100–300 µM), a polyene antibiotic that forms pores permeable for small ions including Na and Cl. Addition of nystatin increased Isc in a biphasic manner (Fig. 1A). Initially, Isc peaked at positive current values (flow of negative charge into lumen) (human: 34.9 ± 10.5 µA/cm2, n = 16; bovine: 109.0 ± 6.0 µA/cm2, n = 98), reversed, and stabilized at negative currents (human: –57.2 ± 13.8 µA/cm2, bovine: –58.2 ± 5.9 µA/cm2). Addition of amiloride (10 µM) had no effect indicating that the apical Na conductance was shunted by nystatin, i.e., the basolateral membrane was rate-limiting for the measured gradient-driven currents. Serosal ouabain (50 µM) was added to eliminate currents generated by the basolateral Na-K-ATPase, which resulted in a further decrease of Isc (Fig. 1A and Table 1).


Figure 1
View larger version (18K):
[in this window]
[in a new window]

  		Fig. 1. Transepithelial measurement of the basolateral Cl conductance (GCl) in Ussing chambers. A: in the presence of a 125-to-5 mM mucosal-to-serosal Cl gradient (i.e., absorptive), addition of nystatin (100 µM) to the mucosal bath induced a biphasic current response. Mucosal addition of amiloride (10 µM) had no effect indicating that the apical membrane was functionally removed. Ouabain (50 µM) was used to block basolateral pump currents generated by the Na-K-ATPase. Cl currents (ICl) across the basolateral membrane returned to 0 after addition of the chloride channel blocker N-phenylanthranilic acid (DPC; 5 mM). B: in Cl-free solutions, nystatin induced a similar current peak but, after addition of ouabain currents, quickly returned to 0. C: addition of forskolin (20 µM) to nystatin-permeabilized preparations further stimulated basolateral Cl currents. Inset compares average values for human and bovine cultures. Responses were not different between human and bovine cultures; P = 0.4 (t-test); number of cultures is given. D: intact, nonpermeabilized cultures (bovine). Note that the direction of the forskolin-stimulated Cl currents was dependent on the direction of the applied Cl gradient. Cl m->s and Cl s->m indicate a mucosal-to-serosal or serosal-to-mucosal Cl gradient across the cultures.

 

View this table:
[in this window]
[in a new window]

  		Table 1. Transepithelial parameters of nystatin-permeabilized human and bovine tracheal sheets

 
In the presence of the applied Cl gradient, Isc is determined by both the basolateral and junctional conductances for Cl. Total transepithelial conductance was 9.9 ± 1.2 mS/cm2 (n = 16, human) and 14.1 ± 0.7 mS/cm2 (n = 74, bovine), as calculated from voltage pulses. Addition of the Cl channel blocker DPC reduced the GCl by 3.4 ± 1.9 mS/cm2 (i.e., 34%, human) and 5.8 ± 0.6 mS/cm2 (41%, bovine) suggesting that a fraction of the measured GCl was mediated by Cl movement across the basolateral membrane. The effects of nystatin, ouabain, and DPC on Isc and Rte are summarized in Table 1.

To further determine the identity of the biphasic nystatin-induced currents, we performed experiments in symmetrical Na-containing but Cl-free solutions (i.e., no gradient). Nystatin induced a similar initial current peak (Fig. 1B) of 76.1 ± 28.5 µA/cm2 (n = 2), and Rte dropped from 2,121 ± 101 {Omega}·cm2 to 737 ± 274 {Omega}·cm2. Addition of ouabain reduced Isc to a value not significantly different from zero (1.3 ± 0.9 µA/cm2, n = 2). In contrast to Cl-containing solutions, no negative currents were measured in absence of Cl. These data suggest that the insertion of nystatin pores in the apical membrane caused 1) initial Na absorption across the apical membrane (the positive current peak) until intracellular [Na] equilibrated with extracellular [Na], which was then followed in the presence of a transepithelial Cl gradient by 2) a sustained Cl current.

Regulation of the basolateral GCl by forskolin. The effect of forskolin on the basolateral GCl was investigated in nystatin-permeabilized tracheal bovine and human cultures (Fig. 1C). Serosal addition of forskolin (20 µM) to nystatin-permeabilized tissues further activated Isc, and addition of DPC blocked a large fraction (Fig. 1C and Table 1). The inset in Fig. 1C compares the effects of forskolin on Isc in permeabilized human and bovine tracheal sheets. Basolateral Isc was stimulated similarly, by 18 ± 5% (n = 16, human) and 23 ± 2% (n = 74, bovine; P = 0.4).

In control experiments, we also tested whether the forskolin-stimulated basolateral GCl is active in tissues not treated with nystatin. This was done by applying both serosal-to-mucosal and mucosal-to-serosal Cl gradients (120-to-5 mM) across intact tissues (Fig. 1D). Figure 1D shows a recording from a bovine tracheal sheet. Under these conditions, a forskolin-regulated and DPC-blockable Cl current is expected to be conducted transcellularly by both the apical and the basolateral GCl. Forskolin activated similar (but reverse) transepithelial Cl currents in the presence of either gradient (serosal-to-mucosal, 11.6 ± 2.3 µA/cm2; mucosal-to-serosal, –10.8 ± 2.5 µA/cm2) and similar conductances (serosal-to-mucosal, 0.6 ± 0.1 mS/cm2; mucosal-to-serosal, 0.6 ± 0.2 mS/cm2, as calculated from the differences of the conductance before and after treatment), and these Cl currents were fully blockable by DPC. These data suggest a GCl present and functional in both the apical and the basolateral cell membrane in intact tracheal sheets.

Effect of Cl channel blockers on basolateral GCl. Next, we tested the effects of different Cl channel blockers in permeabilized, forskolin-stimulated tissues. Individual blockers were related to the effect of DPC (5 mM), which was considered total block of Cl currents and defined as 100% inhibition (6). Figure 2, A–D, illustrates the effects of described Cl channel blockers of the volume-sensitive GCl (sucrose, DIDS, and tamoxifen; Ref. 28). Addition of sucrose to the mucosal chamber dose-dependently and completely inhibited basolateral Cl current, suggesting that all of the measured basolateral GCl is volume-regulated. Half-maximal inhibitory concentration for sucrose was 47 ± 2 mM with a nH of 1.5 ± 0.1 (n = 5, human; Fig. 1B). DIDS inhibited a large fraction of the basolateral GCl, whereas tamoxifen blocked only 11 ± 2% (Fig. 2, E, F, and I).


Figure 2
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 2. Inhibition of the basolateral GCl. Blockers were added to permeabilized, forskolin-stimulated tissues in presence of a mucosal-to-serosal Cl gradient. Bar represents 5 min. A: complete block of basolateral Cl currents by mucosal sucrose (300 mM). Subsequent addition of DPC showed no significant additional effects. B: dose-dependent inhibition of basolateral Cl currents by sucrose. Ki = 47 ± 2 mM; Hill coefficient (nH) = 1.5 ± 0.1. C: effect of serosal DIDS (100 µM, bovine). D: effect of serosal tamoxifen (50 µM, bovine). E and F: block by CdCl2 and ZnCl2 (100 µM, human). Note slow blocking effects. G and H: effects of 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB), flufenamic acid (FFA), and niflumic acid (NFA). Ki and nH were: NPPB, 38 ± 3 µM, 2.2 ± 0.4; FFA, 120 ± 7 µM, 1.5 ± 0.2 ; NFA, 860 ± 88 µM, 0.8 ± 0.07. I: summary of all tested effects of blockers at concentrations to elicit maximal effects. Percent block is compared with block by 5 mM DPC (100%). Blocker effects were similar in human or bovine tissues; t-tests, P = 0.15–1.0; n = 3–10 per bar. Isc, short-circuit current; DNDS, 4,4'-dinitro-2,2'-stilbene-disulfonate disodium salt.

 
Figure 2, E and F, shows the effects of ClC-2-type GCl blockers (CdCl2 and ZnCl2; Ref. 2). Both CdCl2 and ZnCl2 blocked substantial fractions; however, both showed very slow blocking kinetics (Fig. 2, E and F). In addition, we tested glibenclamide as a probe for the CFTR Cl channel, and NPPB, FFA, and NFA were used because of their known effects on the Ca-activated GCl (27). Dose-response relations were investigated for the rapidly acting blockers NPPB, FFA, and NFA (Fig. 2G). The other blockers (DIDS, CdCl2, ZnCl2, and glibenclamide) decreased Cl currents with a slow time course, and therefore dose-response relations were not feasible. NPPB showed the highest affinity with KB = 38 ± 2.9 µM followed by NFA (KB = 120 ± 7 µM) and FFA (KB = 860 ± 9 µM; Fig. 2H).

Figure 2I summarizes the average inhibition of Cl current by the tested blockers in human and bovine tissues. All blockers were used at concentrations to elicit maximal effects. Blocker effects were characterized by rank order (concentration used, % block): DPC (5 mM, 100%) = sucrose (300 mM, 93%) > FFA (1 mM, 83%) = NFA (3 mM, 83%) = glibenclamide (0.5 mM, 82%) > CdCl2 (0.1 mM, 69%) = NPPB (0.2 mM, 68%) = DIDS (0.1 mM, 65%) = ZnCl2 (0.1 mM, 63%) > tamoxifen (0.05 mM, 11%) > DNDS (0.5 mM, 6%) = 0. Sucrose blocked GCl fully (not different from 100, one-sample t-test; P = 0.069). Block by glibenclamide, FFA, and NFA was similar (on average 83.1 ± 2.6%) and significantly larger than NPPB, DIDS, CdCl2, and ZnCl2, which showed similar block (on average 66.5 ± 2.2%; P < 0.0001). Effect by tamoxifen was significant (i.e., different from 0) but smaller than all others (P < 0.0001) excluding DNDS, which showed no significant effect (not different from 0, one-sample t-test; P = 0.072). Effects of blockers were not significantly different in human or bovine cultures (Fig. 2I).

Basolateral GCl is active in intact tracheal sheets. To investigate the role of the basolateral GCl during transepithelial Cl transport, we exploited the DIDS and Cd sensitivity of the basolateral GCl, which distinguishes it from the apical CFTR-mediated GCl. The effects of CdCl2 and DIDS on forskolin-stimulated Cl secretion were determined in symmetrical NaCl solutions in the presence of 10 µM amiloride in intact tissues. Serosal addition of CdCl2 (100 µM) or DIDS (100 µM) increased forskolin-stimulated Isc (Fig. 3). In comparison, addition of CdCl2 or DIDS to the mucosal chamber compartment did not affect Isc (data not shown). The CdCl2- and DIDS-induced currents were completely blocked by addition of 5 mM DPC (apically). These blocker effects indicate that the basolateral GCl is active in intact, transporting epithelia and is a determinant of the magnitude of transepithelial Cl secretion.


Figure 3
View larger version (8K):
[in this window]
[in a new window]

  		Fig. 3. Effects of block of basolateral Cl channels on transepithelial Cl secretion in intact tracheal cultures. A: nonpermeabilized human culture in symmetrical NaCl-Ringer solutions. Na transport was blocked by amiloride (10 µM), and transepithelial Cl secretion was stimulated by 10 µM forskolin. Serosal addition of CdCl2 or DIDS (100 µM each) stimulated Isc. B: average responses to serosal CdCl2 or DIDS of Isc after forskolin stimulation in human and bovine cultures; number of experiments are given in parentheses. Effects in human and bovine cultures were similar (Cd, P = 0.91; DIDS, P = 0.073). Both CdCl2 and DIDS activated currents significantly (P < 0.001).

 
Whole cell Cl conductances in ciliated human airway cells. To further characterize the basolateral GCl, we used both whole cell and cell-attached patch-clamp recordings of single ciliated human airway epithelial cells. The whole cell mode corresponds closely to the nystatin-permeabilized transepithelial preparation. Both methods represent dialyzed preparations, which allow control of the transmembrane ion gradients. Transepithelial experiments were used to localize the conductance to the membrane domain, and whole cell patch-clamp experiments were used to identify Cl conductances by their voltage dependence. Although the sum of both apical and basolateral conductances may be measured in the whole cell patch-clamp mode, we reasoned that, owing to the apical cilia, the apical membrane pole is poorly voltage-clamped, and thus the measured whole cell Cl currents are predominantly conducted across the basolateral pole of the investigated cell.

Whole cell measurements were done within 2 h of isolation of ciliated nasal epithelial cells from 11 healthy volunteers. Cells had a cylindrical shape, and cilia were beating. Although all investigated cells appeared visually similar, there were two functionally distinct cell types of ciliated cells. Forty-three percent (12 out of 28) of the ciliated cells did not express a measurable whole cell Cl conductance even in the presence of forskolin or hypotonic solution. Fifty-seven percent (16 out of 28) of the ciliated cells expressed a significant whole cell Cl conductance. A similar observation was reported in murine nasal epithelial cells, where cells that did not express a Cl conductance were termed type 1 cells, and cells that expressed a measurable Cl conductance were termed type 2 cells (33). In our measurements, type 1 cells were isolated from six individuals who also donated type 2 cells, suggesting that both type 1 and type 2 cells are present in the nasal epithelium of any given individual. Cell membranes of type 1 cells were extremely tight for Cl, and total conductance of the type 1 cells was 9.9 ± 1.4 pS/pF (Cm = 20.9 ± 2.1 pF, n = 12). In 16 type 2 cells, two major and one minor type of whole cell GCl were found distinguished by their voltage-dependent characteristics and the blocker sensitivity.

The most frequently found whole cell Cl conductance was volume-sensitive, DIDS-blockable, and present in 81% of the recordings (Fig. 4). This conductance activated either spontaneously in the whole cell mode or was activated by mild hypotonic treatment (~10% reduced tonicity). Spontaneous current activation was likely caused by cell swelling provoked by the whole cell configuration (44). In the recordings shown in Fig. 4, currents activated spontaneously after establishing the whole cell configuration. Addition of sucrose (Fig. 4A) or DIDS (Fig. 4C) to the bath solution blocked a large portion of the currents. CdCl2 had no effect on this conductance (data not shown). The sucrose- and DIDS-blockable whole cell currents were voltage-dependently activated at both positive and negative potentials resulting in a slightly S-shaped I-V relation (Fig. 4, B, D, and E). The voltage-activating whole cell Cl conductance was termed GVA in this report and was the major whole cell Cl conductance with a current density of GVA = 1,125 ± 288 pS/pF; Cm = 16.7 ± 1.4 pF, n = 13.


Figure 4
View larger version (11K):
[in this window]
[in a new window]

  		Fig. 4. Volume-sensitive and DIDS-sensitive whole cell Cl conductance in human ciliated airway cells. A: dose-dependent block of whole cell Cl current by sucrose. Concentrations of sucrose in mM were added as indicated by arrows. Addition of 100 µM CdCl2 had no effect on this current. Current has been activated in ~10% hypotonic solution before recording. Holding potential was –30 mV pulsed to –20 mV, membrane capacitance (Cm) = 19 pF. During breaks in continuous recording, current-voltage (I-V) relations were obtained. B: I-V step protocols of sucrose-blocked currents. Difference current is shown (before minus after 150 mM sucrose). Excessive noise in this recording resulted from the vibrations caused by the beating cilia of the investigated cell. C: current activated spontaneously after formation of the whole cell configuration. Addition of DIDS (100 µM) blocked whole cell Cl current. I-V relations were recorded before and after addition of DIDS during breaks in recording; holding potential was –40 mV pulsed to –30 mV, Cm = 14 pF. D: I-V step protocols of DIDS-blocked currents. E: steady-state I-V relation was slightly S-shaped, i.e., outwardly rectifying at positive and inwardly rectifying at negative potentials.

 
In 25% of the recordings, an inwardly rectifying and Cd-sensitive whole cell Cl conductance was present, which was termed GIR in this report (Fig. 5). Currents carried by GIR were active immediately after the whole cell formation. In the recording shown in Fig. 5A, current is measured at a baseline voltage of –30 mV and pulsed to –90 mV to monitor the current activation at negative potentials. A large part of the current was blocked by addition of 100 µM CdCl2. The time and voltage dependence of GIR is shown in Fig. 5, B and C. At positive potentials, the steady-state currents were very small; at negative, hyperpolarizing potentials, currents activated. The steady-state I-V relationship of GIR showed strong inward rectification (Fig. 5C). On average, GIR = 750 ± 320 pS/pF, Cm = 17.2 ± 1.7 pF, n = 4.


Figure 5
View larger version (7K):
[in this window]
[in a new window]

  		Fig. 5. Hyperpolarization-activated, inwardly rectifying, and Cd-sensitive whole cell Cl conductance in human ciliated airway cells. A: current recording from a cell held at –30 mV and pulsed every 25 s to –90 mV. During negative pulses, current activation is apparent and was blocked by 100 µM CdCl2. B: I-V step protocol of Cd-blocked current showed current deactivation at positive potentials and current activation at negative potentials. Voltage steps were from +100 to –150 mV. C: steady-state I-V relation of the Cd-blocked current shown in B. Note inward rectification and very small currents at positive potentials.

 
Neither the GVA nor GIR activated when forskolin was added to the bath solution (data not shown). However, in two of 13 stimulations with forskolin, a small distinct whole cell Cl current was activated. Figure 6A shows an example of a forskolin activation of the whole cell Cl current. In this run, GIR and GVA were blocked by previous addition of DIDS and CdCl2. Forskolin-activated Cl currents that were time- and voltage-independent (Fig. 6B), and the I-V relationship was linear (Fig. 6C). The forskolin-activated whole cell Cl conductance was termed GFSK in this report, and averaged GFSK = 96 ± 69 pS/pF, Cm = 13.5 ± 0.5 pF. The voltage dependence of GFSK was clearly distinct from GIR and GVA.


Figure 6
View larger version (7K):
[in this window]
[in a new window]

  		Fig. 6. Forskolin-stimulated whole cell Cl conductance. A: recording from a ciliated human airway cell after treatment with DIDS and CdCl2 (100 µM each). Forskolin (10 µM) activated current. Holding potential was –40 mV pulsed to –30 mV, Cm = 14 pF. B: I-V current step responses. Note time and voltage independence of currents. C: steady-state I-V relations of current activated by forskolin. Slope conductance was 2.75 nS.

 
Single-channel recordings from the basolateral pole of ciliated human airway cells. To identify the properties of the single Cl channels, we performed cell-attached patch-clamp recordings on the basolateral pole of ciliated cells. Cell-attached recordings were done on nasal cells isolated from four individuals. Of 23 successful cell-attached patches, 14 contained Cl channels, and in nine patches no Cl channel activity was recorded even in the presence of forskolin and hypotonic solutions. The Cl channel most frequently found (in 10 of 14 recordings) showed an outwardly rectifying i-V relationship with a g of 59.1 ± 0.6 pS at +100 mV and 21.6 ± 0.3 pS at –100 mV (Fig. 7, A and B). This channel was termed BORC, for basolateral outwardly rectifying Cl channel. On average, the steady-state single-channel Po of BORC was not significantly voltage-dependent (Fig. 7C), and average Po was 0.56 ± 0.03 for all voltages. The time-averaged single-channel current was calculated as i x Po for every potential to estimate the voltage-dependent properties of the average current carried by BORC. The i x Po-to-V relation estimates the voltage-dependent characteristics of the corresponding whole cell current carried by BORC when assuming that the number of channels (N) is not voltage-dependently regulated. The i x Po-to-V relation showed distinct outward rectification (Fig. 7D). BORC was either spontaneously active after formation of the cell-attached mode (in 6 recordings) or activated by hypotonic solution (diluted to 75%, in 4 recordings). A typical example for the activation of BORC by cell swelling and deactivation by 120 mM sucrose is shown in Fig. 7, E and F. In addition, we found that forskolin treatment neither activated silent BORC (in 0 of 4 cases) nor changed Po of already active BORC (before stimulation: Po = 0.40 ± 0.21; after forskolin: Po = 0.52 ± 0.32; n = 4).


Figure 7
View larger version (24K):
[in this window]
[in a new window]

  		Fig. 7. Single-channel recordings of basolateral outwardly rectifying Cl channels (BORC). Cell-attached recording from the basolateral pole of a human ciliated cell. A: typical appearance of single-channel activity at +70 mV (top) and –70 mV (bottom) holding potential. B: single-channel current (i)-V relationship. On average, at +100 mV, single-channel conductance (g) = 59.1 ± 0.6 pS; at –100 mV, g = 21.6 ± 0.3 pS (n = 10). C: mean open probability (Po) was not significantly affected by the holding potential (ANOVA, P = 0.8). Total average was Po = 0.56 ± 0.03. Po was calculated for every voltage by clamping the patch in 25-mV/2-s steps from –100 to +100 mV. D: time-averaged single-channel current, as calculated from i x Po for every holding potential. E: activation of BORC by hypotonic solution. Hypotonic solution (75% tonicity) was added 25 s before start of the shown recording. Holding potential = –70 mV. F: inactivation of channel by hypertonicity. This channel was spontaneously active after seal formation. Sucrose (120 mM) was perfused into the bath as indicated. Holding potential = –60 mV.

 
Basolateral inwardly rectifying Cl channels (termed BIRC) with a chord conductance of g = 10.7 ± 0.23 pS (at –100 mV) and ~3 pS at +75 mV were found in two of 14 cell-attached patches. The recordings were obtained from two different individuals. In both cases, BIRC was spontaneously active after seal formation. One recording is shown in Fig. 8A. At positive potentials, the current amplitude was very small and close to baseline noise. At potentials more negative than –50 mV, single-channel amplitude and activity increased dramatically, resulting in an inwardly rectifying i-V relationship (Fig. 8B). Po of BIRC was highly voltage-dependent (Fig. 8C) and steeply declined from 0.68 ± 0.08 at –100 mV to values close to zero with increasing voltage. The i x Po-to-V relation in Fig. 8D is characterized by a strong inward rectification. At potentials more positive than –50 mV, the time-averaged current was not significantly different from zero. The voltage-dependent properties of BIRC appear similar to GIR measured in the whole cell mode and might be the basis for GIR.


Figure 8
View larger version (15K):
[in this window]
[in a new window]

  		Fig. 8. Single-channel recording of basolateral inwardly rectifying Cl channels (BIRC). Cell-attached recording from the basolateral pole of a human ciliated cell. A: single-channel recording at +80 mV (top) and –80 mV (bottom). This recording was filtered at 65 Hz. B: i-V relation showed inward rectification: at –100 mV, g = 10.7 ± 0.23 pS; at +75 mV, g = 3.2 ± 0.3 pS. C: voltage dependence of Po. D: time-averaged single-channel current, as calculated from i x Po, showed prominent inward rectification. Values between –50 and +75 mV were not significantly different from 0 (one-sample t-tests).

 
Addition of forskolin to the bath activated a channel with characteristics similar to CFTR in two basolateral cell-attached patches. Figure 9A shows the activation of multiple channels in a cell-attached patch. Note that this recording contained multiple basolateral CFTR-like channels (BCFTR) and one large-conductance BORC, which are clearly discernible. Both channel types showed low activity before stimulation. Addition of forskolin significantly increased the activity of BCFTR, but little changed the activity of BORC. Figure 9B shows the Po values for both channel types for the recording shown in Fig. 9A. Po of BCFTR increased substantially over prestimulated levels (Fig. 9, closed circles), whereas Po of BORC remained largely unaffected by stimulation (Fig. 9B, open circles). The I-V relation of BCFTR was linear and averaged 8.1 ± 0.5 pS (Fig. 9C).


Figure 9
View larger version (15K):
[in this window]
[in a new window]

  		Fig. 9. Single-channel recording of a basolateral CFTR-like Cl channel (BCFTR). A: cell-attached recording from the basolateral pole of a human ciliated cell. Initially, a low-conductance and a large-conductance channel show basal activity. Addition of forskolin increased activity of the low-conductance channel. Holding potential = 70 mV; recording was filtered at 50 Hz. B: Po calculated for successive 5-s intervals of the recording in A; Po of BFCR (closed circles) and BORC (open circles) was calculated assuming that this patch contained 1 BORC and 5 BCFTR. C: i-V relation of BCFTR; g = 8.1 ± 0.5 pS.

 
In summary, we have observed three different types of whole cell Cl conductances (Table 2) and three different types of basolateral Cl channels (Table 3). The plots in Fig. 10 display the occurrence and the specific whole cell Cl conductances (Fig. 10A) and the occurrence and calculated specific conductance of each channel type in cell-attached patches (Fig. 10B). In whole cell recordings, GVA was the most frequently observed and largest GCl, followed by GIR and a minor GFSK. The predicted contributions of these to the total GCl, as calculated from the product of the frequency found multiplied by the specific whole cell GCl was GVA (82%) > GIR (16.9%) > GFSK (1.1%). For comparison, in cell-attached patches, BORC was the most frequently observed Cl channel with the largest average single-channel conductance, whereas BIRC and BCFTR occurred less frequently and showed small average single-channel conductances. The estimated contribution of these Cl channels to the total basolateral Cl conductance, as calculated from g x Po x N was BORC (86.5%) > BCFTR (8.3%) > BIRC (5.4%).


View this table:
[in this window]
[in a new window]

  		Table 2. Summary of biophysical characteristics and occurrence of whole cell Cl conductances in human nasal cells

 

View this table:
[in this window]
[in a new window]

  		Table 3. Single-channel characteristics of basolateral Cl channels of human nasal cells

 

Figure 10
View larger version (12K):
[in this window]
[in a new window]

  		Fig. 10. Comparison of the observed whole cell with the cell-attached conductances. A: relative occurrence (closed bars, at –100 mV) and specific conductance (open bars) of the 3 observed whole cell conductances. B: relative occurrence of BORC, BIRC, and BCFTR in cell-attached recordings (closed bars) and their time-averaged conductance (open bars, as calculated from g x Po at –100 mV). GVA, voltage-activating whole cell Cl conductance; GIR, inwardly rectifying and Cd-sensitive whole cell Cl conductance; GFSK, forskolin-activated whole cell Cl conductance.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Most epithelia express a sizable basolateral Cl conductance, which, for example, in the thick ascending limb of the kidney is critical for its normal function of salt absorption (29). Investigations of Cl transport by the airway epithelium has largely focused on the apical Cl conductance owing to its dysfunction in cystic fibrosis. Nevertheless, a basolateral Cl conductance in the airways likely presents a factor to regulate transcellular Cl transport. Previous studies have identified the presence of a basolateral Cl conductance in airways (7, 15, 32, 40, 41), and the current study aimed to determine its functional characteristics. To this end, we combined transepithelial recording from intact and permeabilized preparations with single-channel and whole cell patch-clamp recordings. This comprehensive approach allowed us to relate channel function to epithelial transport and thus determine their physiological role.

The swelling-activated, outwardly rectifying, and DIDS-sensitive basolateral Cl channel. In this study, GVA was the dominant Cl conductance found in whole cell patch-clamp experiments. GVA was characterized by 1) voltage-activation at large positive and negative potentials resulting in an S-shaped steady-state I-V relation, 2) activation by swelling and inactivation by increased osmolality, and 3) block by DIDS (Fig. 4). The voltage dependence of GVA is similar to previous reports of whole cell currents in mouse ciliated nasal cells (33) and in nystatin-permeabilized transepithelial recordings of a basolateral GCl in rat airway epithelium (15). This whole cell conductance may be mediated by BORC, because it was the Cl channel found most frequently (Fig. 10), and 2) activated by swelling and blocked by hyperosmolarity (Fig. 7). However, the outwardly rectifying voltage characteristics of BORC compared with the S-shaped voltage dependence of GVA suggest that either the recording techniques affect the voltage properties (such as differences in ion or ATP concentrations) or GVA represents the sum of more than one channel type. This is currently not clear. Possibly GVA in whole cell recordings is mediated by the sum of activities of both BORC and BIRC, although we have not found that GVA was sensitive to Cd or Zn.

The outwardly rectifying, DIDS-sensitive Cl channel has been investigated for many years in epithelial cells; however, only recently evidence accumulated that this channel may be localized in the basolateral membrane. Using apically nystatin-permeabilized rat or human airway epithelia, Hwang et al. (15) and Szkotak et al. (32) found an outwardly rectifying, DIDS-sensitive basolateral Cl current in transepithelial recordings. For comparison, in intestinal epithelia, outwardly rectifying Cl channels were directly identified by patch-clamping of basolateral membranes of mouse colon (23), mouse jejunum (3), pig enterocytes (25), or pig distal colonic crypts (21). We found BORC in recordings from basolateral membranes of isolated human cells and also found a considerable DIDS-sensitive basolateral GCl in transepithelial recordings of both human and bovine airway cultures suggesting that this channel may be quantitatively the most important contributor to the basolateral GCl of confluent cell sheets.

The hyperpolarization-activated, inwardly rectifying basolateral Cl channel. The whole cell GIR and BIRC in single-channel recordings showed very similar voltage-dependent characteristics suggesting that BIRC found in basolateral membrane patches mediated GIR in whole cell recordings. In addition, the Cd sensitivity of GIR was also found in transepithelial recordings when adding Cd to the serosal side of both nystatin-permeabilized and intact airway cultures. These are strong indications that BIRC and GIR are functional in the basolateral membrane of airways. The basolateral membrane potential of human and bovine tracheal epithelial cells under control conditions was –36 and –74 mV, respectively (35, 39), suggesting that the basolateral membrane potential is a determinant of GIR-mediated Cl currents.

GIR bears remarkable resemblance to currents mediated by ClC-2 when recombinantly expressed in Xenopus oocytes (34) or Sf9 insect cells (45), including strong inward rectification, activation by hyperpolarization, and block by Cd. The molecular expression of ClC-2 in Calu-3 and IB3 airway cell lines has been described (5, 30), and we have found ClC-2 by RT-PCR in primary human airway cultures (data not shown), suggesting that BIRC and GIR are based on the expression of ClC-2. Consistent with the relatively small contribution of GIR to the total basolateral Cl conductance is the reported downregulation of ClC-2 expression in adult compared with fetal tissues (26). Blaisdell et al. (1) and Murray et al. (26) reported that epithelia grown from a mixed population of rat fetal lung epithelial cells expressed ClC-2 apically and not basolaterally using antibody staining and confocal microscopy. In contrast, ClC-2 was found to be expressed at the tight junction complex between adjacent epithelial cells of mouse intestine or CaCo-2 cells (12, 24). Although the different localization might be explained by the different cell types used in these studies, another potential explanation is that the relatively low expression of BIRC in our study is below the sensitivity of antibody detection.

The cAMP-activated, linear Cl channel in the basolateral membrane. In our study, whole cell Cl currents were activated by forskolin in the presence of DIDS and Cd (Fig. 6) suggesting that GFSK is different from the volume-sensitive Cl conductance (9). GFSK was further characterized by its time- and voltage-independent linear currents (Fig. 6), which are functional properties very similar to CFTR-mediated whole cell currents (9, 42). We suggest that the measured whole cell currents are largely basolateral currents because, owing to the series resistance represented by the cilia, the apical membrane can be expected to be poorly space clamped. On the other hand, the observed GFSK was very small, and thus the whole cell recordings are ambiguous as to the location of GFSK. However, in cell-attached recordings, we found evidence for the localization of BCFTR to the basolateral membrane suggesting that a small fraction of the basolateral GCl is mediated by BCFTR. Interestingly, a Cl channel resembling CFTR was also described in the basolateral membrane in renal proximal tubule (31).

Using ciliated mouse nasal epithelial cells, Tarran et al. (33) described whole cell Cl conductances similar to the ones found in our study: a minor time- and voltage-independent Cl conductance (similar to GFSK), a voltage-activating Cl conductance (similar to GVA), and a hyperpolarization-activated Cl conductance (similar to GIR). Although Tarran et al. (33) did not localize these Cl conductances to a membrane pole, we feel that they similarly investigated basolateral conductances because of the effect of space-clamping of ciliated cells in whole cell recordings.

Physiological role of the basolateral Cl conductance during salt and fluid transport in airways. In nystatin-permeabilized tracheal sheets, a large fraction of the basolateral GCl was volume regulated, and forskolin further stimulated Cl currents by ~20% (Fig. 1C). The forskolin-stimulated basolateral GCl was also present in nonpermeabilized, intact epithelia (Fig. 1D). Permeabilization of the apical membrane with nystatin has been shown to be associated with significant cell swelling and activation of volume-sensitive ion conductances (10, 17). Therefore, it was possible that in these experiments, the magnitude of the basolateral Cl conductance was larger than physiologically achieved in intact cells. To test this, we performed additional experiments without nystatin-treatment (Fig. 1D) and compared the effect of forskolin on transepithelial Cl movements in presence of two opposite Cl gradients. In both cases, forskolin activated the transcellular movement of Cl, resulting in a total forskolin-stimulated apical plus basolateral GCl of 0.6 ± 0.2 mS/cm2 in bovine tissues (Fig. 1D). Assuming that the apical GCl was not swelling sensitive and the forskolin-stimulated apical membrane GCl = 1.6 ± 0.3 mS/cm2 (n = 16, bovine; H. Fischer, B. Illek, unpublished results), then the basolateral GCl = 0.95 mS/cm2 in these experiments, which is a value substantially smaller than during nystatin-treatment (5.8 ± 0.6 mS/cm2, bovine). This suggests that the basolateral GCl can be large under certain conditions but physiologically might be smaller and may be a rate-limiting factor during transcellular Cl absorption.

What is the physiological role of the basolateral GCl? Earlier, we have shown that when the electrochemical driving force for Cl is inward across the apical membrane, opening of apical CFTR leads to swelling, activation of the basolateral GCl, and increases in transcellular Cl absorption and transepithelial fluid absorption (35). When the electrochemical gradient for Cl is outward across the apical membrane, tissues will actively secrete Cl. For example, in cAMP-stimulated canine airway epithelium, representative values for the membrane potentials were: apical membrane potential Va = –34 mV; basolateral membrane potential Vb = –62 mV; intracellular Cl activity aCli = 37 mM; and the apical GCl was 4 mS/cm2 (36–38). For these values, the electrochemical driving force for Cl is 13 mV outward across the apical membrane and 38 mV outward across the basolateral membrane. With the electrochemical driving forces for the two membranes and the basolateral GCl of 0.95 mS/cm2 (our study), Cl currents across apical and basolateral membranes in the ratio of ~3:2 are predicted, i.e., a substantial fraction of Cl exits across the basolateral membrane, and Cl secretion is reduced accordingly. If basolateral GCl is increased to the maximum of 5.8 mS/cm2 found in our study, Cl secretion should be inhibited by ~70% if the driving forces for Cl do not change. In fact, an increase in basolateral GCl will further inhibit Cl secretion by depolarizing the cell and by reducing aCli. Although crude, these estimations do suggest that the basolateral GCl can potently influence the degree of Cl secretion.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by National Heart, Lung, and Blood Institute Grants 1P50-HL-60288 and HL-071829, Cystic Fibrosis Foundation Grants 00G0 (B. Illek), 99G0 (H. Fischer), and R613 (W. E. Finkbeiner), and Cystic Fibrosis Research Grant 99-004.


   FOOTNOTES
 

Address for reprint requests and other correspondence: H. Fischer, Children's Hospital Oakland Research Institute, 5700 Martin Luther King, Jr. Way, Oakland, CA 94609 (e-mail: hfischer{at}chori.org)

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.


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Blaisdell CJ, Edmonds RD, Wang XT, Guggino S, Zeitlin PL. pH-regulated chloride secretion in fetal lung epithelia. Am J Physiol Lung Cell Mol Physiol 278: L1248–L1255, 2000.[Abstract/Free Full Text]
  2. Bond TD, Ambikapathy S, Mohammad S, Valverde MA. Osmosensitive Cl currents and their relevance to regulatory volume decrease in human intestinal T84 cells: outwardly vs. inwardly rectifying currents. J Physiol 511: 45–54, 1998.[Abstract/Free Full Text]
  3. Butt AG, Hamilton KL. Ion channels in isolated mouse jejunal crypts. Pflügers Arch 435: 528–538, 1998.[CrossRef][Web of Science][Medline]
  4. Chang D, Dawson DC. Demonstration of calcium-activated basolateral K+ and Cl– conductances. J Gen Physiol 92: 281–306, 1988.[Abstract/Free Full Text]
  5. Cuppoletti J, Tewari KP, Sherry AM, Kupert EY, Malinowska DH. ClC-2 Cl– channels in human lung epithelia: activation by arachidonic acid, amidation, and acid-activated omeprazole. Am J Physiol Cell Physiol 281: C46–C54, 2001.[Abstract/Free Full Text]
  6. Di Stefano A, Wittner M, Schlatter E, Lang HJ, Englert H, Greger R. Diphenylamine-2-carboxylate, a blocker of the Cl–-conductive pathway in Cl–-transporting epithelia. Pflügers Arch 405, Suppl 1: S95–S100, 1985.[CrossRef][Web of Science][Medline]
  7. Duta V, Szkotak A, Nahirney D, Duszyk M. The role of bestrophin in airway epithelial ion transport. FEBS Lett 577: 551–554, 2004.[CrossRef][Web of Science][Medline]
  8. Fischer H, Machen TE. CFTR displays voltage dependence and two gating modes during stimulation. J Gen Physiol 104: 541–566, 1994.[Abstract/Free Full Text]
  9. Foskett JK. ClC and CFTR chloride channel gating. Annu Rev Physiol 60: 689–717, 1998.[CrossRef][Web of Science][Medline]
  10. Germann WJ, Ernst SA, Dawson DC. Resting and osmotically induced basolateral K conductances in turtle colon. J Gen Physiol 88: 253–274, 1986.[Abstract/Free Full Text]
  11. Grygorczyk R, Bridges MA. Whole-cell chloride conductances in cultured brushed human nasal epithelial cells. Can J Physiol Pharmacol 70: 1134–1141, 1992.[Web of Science][Medline]
  12. Gyomorey K, Yeger H, Ackerley C, Garami E, Bear CE. Expression of the chloride channel ClC-2 in the murine small intestine epithelium. Am J Physiol Cell Physiol 279: C1787–C1794, 2000.[Abstract/Free Full Text]
  13. Halm DR. Identifying swelling-activated channels from ion selectivity patterns. J Gen Physiol 112: 369–371, 1998.[Free Full Text]
  14. Hanrahan JW, Alles WP, Lewis SA. Single anion-selective channels in the basolateral membrane of a mammalian tight epithelium. Proc Natl Acad Sci USA: 7791–7795, 1985.
  15. Hwang TH, Lee HJ, Lee NK, Choi YC. Evidence that basolateral but not apical membrane localization of outwardly rectifying depolarization-induced Cl– channel in airway epithelia. J Membr Biol 176: 217–221, 2000.[CrossRef][Web of Science][Medline]
  16. Illek B, Fischer H. Flavonoids stimulate Cl conductance of human airway epithelium in vitro and in vivo. Am J Physiol Lung Cell Mol Physiol 275: L902–L910, 1998.[Abstract/Free Full Text]
  17. Illek B, Fischer H, Kreusel KM, Hegel U, Clauss W. Volume-sensitive basolateral K+ channels in HT-29/B6 cells: block by lidocaine, quinidine, NPPB, and Ba2+. Am J Physiol Cell Physiol 263: C674–C683, 1992.[Abstract/Free Full Text]
  18. Illek B, Tam AWK, Fischer H, Machen TE. Anion selectivity of apical membrane conductance of Calu 3 human airway epithelia. Pflügers Arch 437: 812–822, 1999.[CrossRef][Web of Science][Medline]
  19. Jiang C, Finkbeiner WE, Widdicombe JH, McCray PB, Miller SS. Altered fluid transport across airway epithelium in cystic fibrosis. Science 262: 424–427, 1993.[Abstract/Free Full Text]
  20. Kondo M, Finkbeiner WE, Widdicombe JH. Cultures of bovine tracheal epithelium with differentiated ultrastructure and ion transport. In Vitro Cell Dev Biol 29: 19–24, 1992.[CrossRef]
  21. Li Y, Halm ST, Halm DR. Secretory activation of basolateral membrane Cl– channels in guinea pig distal colonic crypts. Am J Physiol Cell Physiol 284: C918–C933, 2003.[Abstract/Free Full Text]
  22. Marcus DC, Shunji T, Wangemann P. Two types of chloride channels in the basolateral membrane of vestibular dark cells. Hear Res 69: 124–132, 1993.[CrossRef][Web of Science][Medline]
  23. Mignen O, Egee S, Liberge M, Harvey BJ. Basolateral outward rectifier chloride channel in isolated crypts of mouse colon. Am J Physiol Gastrointest Liver Physiol 279: G277–G287, 2000.[Abstract/Free Full Text]
  24. Mohammad-Panah R, Gyomorey K, Rommens J, Choudhury M, Li C, Wang Y, Bear CE. ClC-2 contributes to native chloride secretion by a human intestinal cell line, Caco-2. J Biol Chem 276: 8306–8313, 2001.[Abstract/Free Full Text]
  25. Monaghan AS, Mintenig GM, Sepulveda FV. Outwardly rectifying Cl– channel in guinea pig small intestinal villus enterocytes: effect of inhibitors. Am J Physiol Gastrointest Liver Physiol 273: G1141–G1152, 1997.[Abstract/Free Full Text]
  26. Murray CB, Chu S, Zeitlin PL. Gestational and tissue-specific regulation of ClC-2 chloride channel expression. Am J Physiol Lung Cell Mol Physiol 271: L829–L837, 1996.[Abstract/Free Full Text]
  27. Nilius B, Prenen J, Voets T, Van den Bremt K, Eggermont J, Droogmans G. Kinetic and pharmacological properties of the calcium-activated chloride-current in macrovascular endothelial cells. Cell Calcium 22: 53–63, 1997.[CrossRef][Web of Science][Medline]
  28. Nilius B, Sehrer J, Viana F, De Greef C, Raeymaekers L, Eggermont J, Droogmans G. Volume-activated Cl– currents in different mammalian non-excitable cell types. Pflügers Arch 428: 364–371, 1994.[CrossRef][Web of Science][Medline]
  29. Paulais M, Teulon J. cAMP-activated chloride channel in the basolateral membrane of the thick ascending limb of the mouse kidney. J Membr Biol 113: 253–260, 1990.[CrossRef][Web of Science][Medline]
  30. Schwiebert EM, Cid-Soto LP, Stafford D, Carter M, Blaisdell CJ, Zeitlin PL, Guggino WB, Cutting GR. Analysis of ClC-2 channels as an alternative pathway for chloride conduction in cystic fibrosis airway cells. Proc Natl Acad Sci USA 95: 3879–3884, 1998.[Abstract/Free Full Text]
  31. Segal AS, Geibel J, Boulpaep EL. A chloride channel resembling CFTR on the basolateral membrane of rabbit proximal tubule (Abstract). J Am Soc Nephrol 4: 879, 1993.
  32. Szkotak AJ, Man SFP, Duszyk M. The role of the basolateral outwardly rectifying chloride channel in human airway epithelial anion secretion. Am J Respir Cell Mol Biol 29: 710–720, 2003.[Abstract/Free Full Text]
  33. Tarran R, Gray MA, Evans MJ, Colledge WH, Ratcliff R, Argent BE. Basal chloride currents in murine airway epithelial cells: modulation by CFTR. Am J Physiol Cell Physiol 274: C904–C913, 1998.[Abstract/Free Full Text]
  34. Thiemann A, Grunder S, Pusch M, Jentsch TJ. A chloride channel expressed in epithelial and non-epithelial cells. Nature 356: 57–60, 1992.[CrossRef][Medline]
  35. Uyekubo S, Fischer H, Manishikis C, Illek B, Miller SS, Widdicombe JH. Cyclic AMP-dependent absorption of salt and water across airways. Am J Physiol Lung Cell Mol Physiol 275: L1–L9, 1998.[Abstract/Free Full Text]
  36. Welsh M, Smith P, Frizzell R. Chloride secretion by canine tracheal epithelium: II. The cellular electrical potential profile. J Membr Biol 70: 227–238, 1982.[CrossRef][Web of Science][Medline]
  37. Welsh M, Smith P, Frizzell R. Chloride secretion by canine tracheal epithelium: III. Membrane resistances and electromotive forces. J Membr Biol 71: 209–218, 1983.[CrossRef][Web of Science][Medline]
  38. Welsh MJ. Intracellular chloride activities in canine tracheal epithelium. Direct evidence for sodium-coupled intracellular chloride accumulation in a chloride-secreting epithelium. J Clin Invest 71: 1392–1401, 1983.[Web of Science][Medline]
  39. Willumsen NJ, Boucher RC. Shunt resistance and ion permeabilities in normal and cystic fibrosis airway epithelia. Am J Physiol Cell Physiol 256: C1054–C1063, 1989.[Abstract/Free Full Text]
  40. Willumsen NJ, Davis CW, Boucher RC. Cellular Cl– transport in cultured cystic fibrosis airway epithelium. Am J Physiol Cell Physiol 256: C1045–C1053, 1989.[Abstract/Free Full Text]
  41. Willumsen NJ, Davis CW, Boucher RC. Intracellular Cl– activity and cellular Cl– pathways in cultured human airway epithelium. Am J Physiol Cell Physiol 256: C1033–C1044, 1989.[Abstract/Free Full Text]
  42. Winpenny JP, McAlroy HL, Gray MA, Argent BE. Protein kinase C regulates the magnitude and stability of CFTR currents in pancreatic duct cells. Am J Physiol Cell Physiol 268: C823–C828, 1995.[Abstract/Free Full Text]
  43. Wladkowski SL, Lin W, McPheeters M, Kinnamon SC, Mierson S. A basolateral Cl conductance in rat lingual epithelium. J Membr Biol 164: 91–101, 1998.[CrossRef][Web of Science][Medline]
  44. Worrell RT, Butt AG, Cliff WH, Frizzell RA. A volume-sensitive choride conductance in human colonic cell line T84. Am J Physiol Cell Physiol 256: C1111–C1119, 1989.[Abstract/Free Full Text]
  45. Xiong H, Li C, Garami E, Wang Y, Ramjeesingh M, Galley K, Bear CE. ClC-2 activation modulates regulatory volume decrease. J Membr Biol 167: 215–221, 1999.[CrossRef][Web of Science][Medline]
  46. Yamaya M, Finkbeiner WE, Chun SY, Widdicombe JH. Differentiated structure and function of cultures from human tracheal epithelium. Am J Physiol Lung Cell Mol Physiol 262: L713–L724, 1992.[Abstract/Free Full Text]
  47. Zabner J, Smith JJ, Karp PH, Widdicombe JH, Welsh MJ. Loss of CFTR chloride channels alters salt absorption by cystic fibrosis airway epithelia in vitro. Mol Cell 2: 397–403, 1998.[CrossRef][Web of Science][Medline]



This article has been cited by other articles:


Home page
 JGPHome page
R. J. Lee, J. M. Harlow, M. P. Limberis, J. M. Wilson, and J. K. Foskett
HCO3- Secretion by Murine Nasal Submucosal Gland Serous Acinar Cells during Ca2+-stimulated Fluid Secretion
J. Gen. Physiol., July 1, 2008; 132(1): 161 - 183.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
J. Brillault, T. I. Lam, J. M. Rutkowsky, S. Foroutan, and M. E. O'Donnell
Hypoxia effects on cell volume and ion uptake of cerebral microvascular endothelial cells
Am J Physiol Cell Physiol, January 1, 2008; 294(1): C88 - C96.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
292/6/L1432    most recent
00032.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (2)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Fischer, H.
Right arrow Articles by Widdicombe, J. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Fischer, H.
Right arrow Articles by Widdicombe, J. H.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.