Acid secretion and proton conductance in human airway epithelium

Horst Fischer, Jonathan H. Widdicombe, and Beate Illek

Children's Hospital Oakland Research Institute, Oakland, California 94609


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Acid secretion and proton conductive pathways across primary human airway surface epithelial cultures were investigated with the pH stat method in Ussing chambers and by single cell patch clamping. Cultures showed a basal proton secretion of 0.17 ± 0.04 µmol · h-1 · cm-2, and mucosal pH equilibrated at 6.85 ± 0.26. Addition of histamine or ATP to the mucosal medium increased proton secretion by 0.27 ± 0.09 and 0.24 ± 0.09 µmol · h-1 · cm-2, respectively. Addition of mast cells to the mucosal medium of airway cultures similarly activated proton secretion. Stimulated proton secretion was similar in cultures bathed mucosally with either NaCl Ringer or ion-free mannitol solutions. Proton secretion was potently blocked by mucosal ZnCl2 and was unaffected by mucosal bafilomycin A1, Sch-28080, or ouabain. Mucosal amiloride blocked proton secretion in tissues that showed large amiloride-sensitive potentials. Proton secretion was sensitive to the application of transepithelial current and showed outward rectification. In whole cell patch-clamp recordings a strongly outward-rectifying, zinc-sensitive, depolarization-activated proton conductance was identified with an average chord conductance of 9.2 ± 3.8 pS/pF (at 0 mV and a pH 5.3-to-pH 7.3 gradient). We suggest that inflammatory processes activate proton secretion by the airway epithelium and acidify the airway surface liquid.

primary airway cultures; JME airway cells; pH stat; patch clamp; Ussing chamber


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

THE MUCOSA OF THE AIRWAY SURFACE epithelium is lined with a thin layer of fluid called the airway surface liquid (ASL). The composition of the ASL affects its physiological functions, the most important of which are removal of inhaled particles and antimicrobial activity (36). Active transport of Na+ and Cl- by the airway epithelium have been well characterized and recognized as critical determinants of ASL composition and depth. In contrast, the regulation of the H+ concentration in the ASL has received little attention. Acidic luminal pH has been shown to inhibit ciliary beating (4) and to cause bronchoconstriction (1), cough (39), loosening of the epithelial cells from one another (13), and detachment from the basement membrane (17). Acidic ASL has been implicated in airway diseases such as asthma (19, 29) and cystic fibrosis (CF) (33). Although the airway epithelium has been shown to secrete HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>, normal ASL in ferret trachea was previously reported as slightly acidic (pH 6.85) compared with plasma (26). In human primary airway cultures a pH of 6.9 was recently found with the use of either pH-selective microelectrodes (5) or a pH-sensitive fluorophore (22).

The concentration of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> is one determinant of the pH of ASL, and active secretion of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> across primary airway cultures or cell lines has been demonstrated (28, 33), presumably reflecting the HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> permeability of the apical CF transmembrane conductance regulator (CFTR) Cl- channel (31). Jayaraman et al. (22) estimated a HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> concentration of 8 mM in the ASL from their measurements of pH and a partial pressure of CO2 of 5 kPa. With that HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> concentration and with an average CO2 level in the airways in vivo of 2.2 kPa, the ASL is predicted to be slightly alkaline (pH ~7.3), which is in contrast to the reported relative acidity of ASL. Thus we hypothesized that the airway epithelium expresses a regulated mechanism to secrete H+ into the ASL. Because the volume of the ASL is small (~1 µl/cm2) it can be expected to be rapidly acidified by cellular H+ transport. Recently, Hunt et al. (19) found that the exhaled breath of asthmatic patients during an attack is markedly acidic (pH 5.2). We hypothesized that during an inflammatory challenge the airway epithelium secretes increased amounts of acid into the ASL. In this report we measured acid secretion by human airway epithelial cultures in vitro and investigated its mechanism and regulation. We conclude that mucosal histamine, ATP, or mast cells activate epithelial acid secretion that is mediated mainly by an electrogenic apical proton conductance.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell cultures. Human tracheal primary cultures were isolated and cultured as previously described (40). In brief, strips of epithelium were removed from the underlying tissues and treated with protease overnight. The resulting isolated, dispersed cells were plated on permeable filter supports (Snapwell, 0.4-µm pore size, 1-cm2 area; Corning Costar, Cambridge, MA) precoated with human placental collagen (15 µg/cm2) at a density of ~106 cells/cm2. Cells were grown in DMEM-F12 culture medium supplemented with 2% Ultroser G (Biotechnics, Paris, France) and antibiotics. Tracheal sheets were grown to confluence in an air-liquid interface in a tissue culture incubator gassed with 5% CO2 and air. The cystic fibrosis JME/CF15 airway cell line (23) was cultured in a DMEM-F-12 mixture supplemented with 10% fetal bovine serum and (per ml) 5 µg of insulin, 0.5 µg of hydrocortisone, 10 ng of epidermal growth factor, 5 ng of transferrin, 1.3 ng of triiodothyroxine, 43 ng of adenine, and 1 µg of epinephrine. For patch clamping, cells were seeded at low density on cover glasses. The human mast cell line HMC-1 was kindly provided by Dr. J. H. Butterfield and was cultured as described previously (2).

Quantification of proton secretion. Proton secretion was measured with the pH stat titration method in an Ussing chamber (14). Cultures (1-cm2 exposed area) were mounted in an Ussing chamber (Physiologic Instruments, San Diego, CA) and, unless otherwise described, bathed serosally with HEPES-buffered solution and mucosally with buffer-free solution (3 ml each). Solutions were constantly gassed with oxygen and were nominally free of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2. The top of the open chamber was gassed with nitrogen to prevent air CO2 from entering the solutions. Standard NaCl Ringer solutions contained (in mM) 140 NaCl, 2 KCl, 15 glucose, 2 CaCl2, and 1 MgCl2 (mucosal) or 140 NaCl, 2 KCl, 5 glucose, 10 HEPES, 2 CaCl2, and 1 MgCl2 (serosal), adjusted to pH 7.3 with N-methyl-D-glucamine (NMDG). In some experiments the serosal solution was HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2 buffered, containing (in mM) 120 NaCl, 25 NaHCO3, 2 KCl, 5 glucose, 2 CaCl2, and 1 MgCl2, gassed with 5% CO2-95% O2. The pH of the mucosal solution was continuously measured, amplified, and recorded on a computer through a 12-bit analog-to-digital converter. Mucosal pH was manually titrated to a target value of pH 7.3 with 10 mM NaOH made from certified standards. Volumes on the order of 0.5-2.0 µl were added with a precision pipette (Eppendorf Ultra Micro 2.5; Brinkmann Instruments). From the added volume and the time between identical pH readings in the recording the rate of H+ secretion (JH) was determined. Rates are expressed in micromoles per hour per square centimeter, and positive rates refer to acidification of the mucosal medium. To test the quality of our measurements and to identify the cultures as the source of the measured mucosal acidity we did control experiments without cells for all conditions. No component of our recording system released significant acid equivalents, and pH was stable over the course of several hours. Tissues were investigated under current-clamped conditions. Unless otherwise described, current was clamped to zero. Transepithelial potential (Vt, referenced to the serosal side) was continuously recorded, and transepithelial resistance (Rt) was determined from 20-µA pulses. Positive transepithelial currents refer to cation secretion into the mucosal bath. Measurements were done at 35-37°C.

Whole cell patch-clamp recordings. Cells were whole cell patch-clamped after 2 days in culture as previously described (20) on a temperature-controlled stage (35-37°C) of an inverted microscope. Cells were bathed in (in mM) 100 HEPES, 85 gluconic acid, 100 NMDG, 2 Ca gluconate, 1 Mg gluconate, and 10 glucose, pH 7.3. The pipette was filled with (in mM) 100 2-(N-morpholino)ethanesulfonic acid (MES), 81.6 gluconic acid, 70 NMDG, 10 NMDG-EGTA, 1 glucose, 1 MgCl2, 3.3 Mg-ATP, and 0.07 Li-GTP, pH 5.3. The bath electrode was made with the pipette-filling solution (but ATP/GTP free) and a 3% agar bridge. Junction potentials were measured, zeroed, and carefully observed for stability. With these solutions, pipette resistance was ~5 MOmega . Only seals >50 GOmega were used for recordings. The access resistance (Ra) and the cell membrane capacitance (Cm) were determined by fitting the current transients caused by a 10-mV voltage pulse with a single exponential. Measured Ra was 15.4 ± 1.8 MOmega (n = 13), and Cm was 29.1 ± 4.3 pF. Current-voltage step protocols from -80 mV to +60 mV were applied, and resulting step currents were recorded. Whole cell conductance was calculated as the chord conductance at 0 mV. When the specific membrane conductance (Gm in pS/pF) was calculated, whole cell conductance was corrected for Ra and normalized to Cm.

Drugs. Stock solutions of histamine (free base, 10 mM), ATP (Na salt, 50 mM), and ZnCl2 (1 mM) were prepared in NaCl Ringer solution, and pH was adjusted to 7.3. Amiloride stock was prepared at 10 mM in water, and pH was adjusted to 7.3. 3-(Cyanomethyl)-2-methyl-8-(phenylmethoxy)-imidazo- [1,2a]-pyridine (Sch-28080) was kindly provided by Dr. T. E. Machen (Univ. of California, Berkeley) and prepared as a 5 mM stock in ethanol. Bafilomycin A1 (500 µM) and ouabain (100 mM) were prepared as stocks in dimethyl sulfoxide.

Statistics. Data are given as original values or as means ± SE; n is the number of cultures tested. Unpaired t-tests were used to test for statistical difference (P < 0.05) between means, and one-sample t-tests were used to test whether responses were significantly different from zero.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Proton secretion across human airway cultures was investigated in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2-free solutions in Ussing chambers. At a mucosal pH of 7.3 all tested cultures acidified the mucosal medium. In NaCl solutions the basal rate of acidification was JH = 0.17 ± 0.04 µmol · h-1 · cm-2 (n = 26). Basal Vt was -15.6 ± 4.1 mV, and Rt was 1,016 ± 137 Omega  · cm2. Figure 1 shows an example of a measurement of mucosal pH and determination of the basal rate of acidification. Initially, the pH was titrated two times with NaOH to ~7.3 to calculate the rate of acidification. When no NaOH was added, the pH reached an equilibrium at 6.85 ± 0.26 (n = 5) after 41 ± 9 min.


View larger version (8K):
[in this window]
[in a new window]
 
Fig. 1.   Recording of mucosal pH of a human airway culture. Initially pH was titrated to 7.3 to calculate the rate of acid secretion by the tissue. The mucosal bath was then allowed to acidify until an equilibrium was reached. In this experiment the equilibrium pH was 6.98.

Addition of histamine or ATP (100 µM each) to the mucosal bath potently stimulated H+ secretion (Fig. 2, A and B). Both agents typically caused a fast initial rise followed by a slow decay of JH. Figure 2C shows the average peak effects of mucosal addition of histamine and ATP. In contrast, when histamine or ATP was added to the serosal bath, JH was unchanged (not shown). We also tested 50 µM serotonin and 20 µM forskolin, which showed no significant effects on JH [serotonin, +0.04 ± 0.05 µmol · h-1 · cm-2 (n = 3); forskolin, +0.01 ± 0.10 µmol · h-1 · cm-2 (n = 3)].


View larger version (26K):
[in this window]
[in a new window]
 
Fig. 2.   Rates of mucosal acidification in human airway cultures. A and B: effects of histamine (Hist) and ATP (100 µM each) on H+ secretion when added to the mucosal medium. C: average activated rates of H+ secretion by 100 µM histamine and 100 µM ATP. Peak rates were averaged. Nos. in parentheses are nos. of cultures tested. Both histamine and ATP significantly reduced transepithelial resistance (Rt) to, on average, 82.2 ± 5.1% (histamine) and 68.2 ± 4.6% (ATP) of the values before agonist addition. D: effects of 5 µM Sch-28080 (Sch), 100 nM bafilomycin A1 (BafA1), and 400 µM amiloride (Ami). All blockers were added to the mucosal medium after tissues were stimulated with histamine and ATP (100 µM each). E: effects of 200 µM ZnCl2 (Zn) on stimulated H+ secretion. F: average effects of blockers. Effects of bafilomycin A1, Sch-28080, ouabain (Ouab), and amiloride were not different from zero (by 1-sample t-test). Amiloride blocked H+ secretion in 4 cultures and showed no effect in 7 others. These drugs showed no significant effects on Rt (in % of predrug addition: bafilomycin A1, 103 ± 3; Sch-28080, 92 ± 4; ouabain, 95 ± 4; amiloride, 108 ± 16). Block of H+ secretion by ZnCl2 was significant (P = 0.032). ZnCl2 also increased Rt significantly to 114 ± 4% (P = 0.028) of values recorded before ZnCl2 addition. Nos. in parentheses are nos. of cultures tested. G: effects of 100 µM mucosal ouabain followed by 200 µM ZnCl2 on stimulated H+ secretion. Effect of ouabain was small and on average not significant. H: stimulation of H+ secretion in absence of mucosal ions. Airway culture was bathed serosally with NaCl Ringer solution and mucosally with ion-free 300 mM mannitol. ATP (100 µM) was added to the mucosal bath. y-Axes in A, B, D, E, G, and H are absolute values; y-axes of C and F are differences. Time bar is the same for all records.

To identify the mechanism(s) in the apical membrane responsible for the H+ secretion we tested known blockers of H+ transporters. Cultures were stimulated with a combination of histamine and ATP, which we found to result in more sustained and stable responses compared with application of the single agonists. Combined treatment resulted in peak responses of 0.59 ± 0.06 µmol · h-1 · cm-2 (n = 13). We tested the effects of the following blockers: 5 µM Sch-28080 to probe for the gastric-type K+-H+-ATPase, 100 µM ouabain to probe for the non-gastric-type K+-H+-ATPase, 100 nM bafilomycin A1 to probe for the V-type H+-ATPase, 400 µM amiloride to probe for the Na+/H+ exchanger, and 200 µM ZnCl2 to block H+ channels. All blockers were added to the mucosal side. Figure 2, D, E, and G, shows typical blocker experiments, and Fig. 2F summarizes the average effects of blockers. Both bafilomycin A1 and Sch-28080 are highly specific and selective blockers. Neither showed significant effects on JH (effects not different from zero, 1-sample t-tests), indicating that the V-type H+-ATPase and the gastric-type K+-H+-ATPase do not significantly contribute to H+ secretion across airways. Similarly, mucosal ouabain (Fig. 2, G and F) showed very small effects that were not significantly different from zero, indicating that the non-gastric-type K+-H+-ATPase contributes little to H+ secretion.

Addition of amiloride to the mucosal bath had variable effects on H+ secretion. In 4 of 11 cultures tested JH was transiently inhibited by -0.17 ± 0.07 µmol · h-1 · cm-2 (peak inhibition), corresponding to an inhibition of 34% of total H+ secretion. Figure 2D shows an example. In the other seven cultures tested amiloride had no significant effect (0.02 ± 0.03 µmol · h-1 · cm-2). In Fig. 2F the total average effect of amiloride is given. Interestingly, the cultures that showed an amiloride-sensitive JH also expressed a larger amiloride-sensitive Vt. In these cultures amiloride blocked 76 ± 19% of Vt, whereas in the cultures that did not show an amiloride-sensitive JH, Vt was blocked by only 17 ± 8%. This suggested that the effects of amiloride on JH were likely not mediated by an apical Na+/H+ exchanger. However, the data are consistent with the notion that the activity of apical Na+ channels affected JH (see DISCUSSION).

ZnCl2 (200 µM) added to the mucosal bath caused sustained block of JH (Fig. 2, E and G). ZnCl2 was the only blocker used that consistently blocked a large fraction of JH. In nine of nine cultures tested, 200 µM ZnCl2 blocked on average 70 ± 8.8% of JH. These data suggest that a zinc-sensitive H+ conductance is the major mechanism for the transepithelial JH in airway cultures.

In addition, we measured H+ secretion in cultures that were incubated on the mucosal side with nominally ion-free solution (300 mM mannitol) to determine the ion dependence of the apical mechanism. Serosal solution was standard NaCl-HEPES Ringer solution. Under these conditions the basal rate of acidification was 0.07 ± 0.03 µmol · h-1 · cm-2 (n = 3). Addition of ATP to the mucosal bath resulted in an increase of JH of 0.28 ± 0.02 µmol · h-1 · cm-2. An example is shown in Fig. 2H. Addition of histamine resulted in peak increases of JH of 0.18 ± 0.03 µmol · h-1 · cm-2 (n = 2), and histamine plus ATP resulted in increases of 0.68 ± 0.08 µmol · h-1 · cm-2 (n = 2). The rates measured in ion-free solutions were not different from the rates obtained in NaCl Ringer solutions.

The zinc sensitivity and the ion independence of stimulated H+ secretion indicated an apical H+ conductance. Therefore, JH should be dependent on the electrochemical driving force across the apical membrane. To test this hypothesis transepithelial currents were passed across the epithelium. Airway cultures were bathed in NaCl Ringer, and transepithelial current was clamped for 10 min sequentially to -200, 0, or +200 µA/cm2 before and after ATP stimulation. Figure 3 shows JH measured at different currents. It should be noted that JH was positive at all clamped currents. Positive currents (which depolarize the apical membrane) significantly increased JH . The inhibition of JH by negative currents (which hyperpolarize the apical membrane) was less than the observed stimulation by positive currents. This relation suggested outward rectification of H+ currents with respect to the apical membrane potential.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 3.   Proton secretion is affected by injected current. Constant currents were passed across the epithelium, and H+ secretion into the mucosal compartment was measured. Data are from 3 tissues before (open circle ) and after () ATP activation. For some points, SE bars are smaller than symbols.

To measure H+ currents directly, single JME airway cells were whole cell patch-clamped under conditions selective for H+ currents (see METHODS). For the patch-clamp experiments JME cells were used in preference to primary ciliated airway cells because of 1) the difficulty in obtaining high-resistance seals on ciliated cells and 2) the poor space clamping of the apical membrane of ciliated cells. H+ currents were measured selectively under conditions adapted from the measurement of H+ currents in alveolar cells (3), and cells were prestimulated with 100 µM ATP in the bath. H+ currents were identified by 1) reversal potential, 2) voltage-dependent activation, and 3) sensitivity to zinc. Figure 4 shows typical whole cell recordings. In the presence of a pH 5.3-to-pH 7.3 proton gradient from cell to bath H+ currents showed a negative reversal potential of -72 ± 6.1 mV (n = 13). When membrane potential was stepped to depolarizing potentials, currents showed slow activation and strong outward rectification (Fig. 4, A and B), which are typical characteristics of H+ currents (3). On average, we found a steady-state H+ conductance of 9.2 ± 3.8 pS/pF (n = 13, chord conductance at 0 mV, with an average cell capacitance of 29 ± 4 pF). Figure 4C shows a whole cell patch-clamp experiment in which the effect of ZnCl2 was tested. The membrane potential was clamped to -80 mV and pulsed to +20 mV every 10 s to continuously monitor the voltage activation of the H+ current as an identifier. Before addition of ZnCl2, large currents activated during the depolarizing pulses. Addition of ZnCl2 to the bath readily blocked the voltage-activated positive (outward) currents. Figure 4D shows details of the voltage-activated currents before and after addition of ZnCl2.


View larger version (26K):
[in this window]
[in a new window]
 
Fig. 4.   Whole cell proton currents in JME airway cells. A: current responses to voltage steps from -80 mV to +60 mV in 20-mV steps. Holding potential between steps was -60 mV. Note that H+ currents were small at negative potentials. Outward currents at positive potentials were voltage-dependently activated. B: current-voltage relations. Steady-state currents were outwardly rectifying. Reversal potential was -62 mV in this recording, indicative of H+ currents. Composition of solutions was chosen such that no other ion species could generate a negative reversal potential. C: continuous current recording at a holding potential of -80 mV, pulsed every 10 s for 5 s to +20 mV. A large part of the current elicited by a depolarizing voltage pulse was blocked by 200 µM ZnCl2. D: 2 typical pulses selected from the recording in A. Note the slow depolarization-induced activation of current (control) and its block by Zn2+ (Zn). Inset: applied pH gradient in whole cell patch-clamp mode.

In vivo the main source for mucosal histamine in the airways is probably mucosal mast cells. To test the direct effect of mast cells on H+ secretion by the airway epithelium we used the human mast cell line HMC-1 (2). HMC-1 cells constitutively degranulate and release histamine and other factors (2). HMC-1 cells were suspended in unbuffered NaCl Ringer, the pH was adjusted to 7.3, and a cell suspension containing 1.5 × 106 HMC-1 cells was added to the mucosal compartment. Figure 5A shows a typical experiment. Addition of HMC-1 cells resulted in a large activation of H+ secretion. On average, JH values of 0.87 ± 0.33 µmol · h-1 · cm-2 (n = 3; peak value) were stimulated, which is a significantly larger response than treatment with histamine (0.27 ± 0.09 µmol · h-1 · cm-2; Fig. 2C). The increased acidification may have been caused by another factor released by the mast cells or by H+ release by the mast cells. This latter possibility was tested by adding HMC-1 cells to experiments without airway cultures (Fig. 5B). When added, 1.5 × 106 HMC-1 cells showed a brief release of acid with a rate of 0.56 ± 0.028 µmol/h (n = 4 experimental runs). Within <10 min JH from HMC-1 cells returned to zero, indicating that parts of the initial peak, but not the continuous acid secretion by airway epithelium, was caused by a brief acid release from HMC-1 cells.


View larger version (11K):
[in this window]
[in a new window]
 
Fig. 5.   Mucosal mast cells activate H+ secretion. A: airway culture was bathed in NaCl Ringer solution. HMC-1 mast cells (1.5 × 106) were added to the mucosal medium where indicated by arrow. B: control experiment without airway epithelial cells present. HMC-1 mast cells (1.5 × 106) were added to the cell-free insert where indicated by arrow.

We next performed measurements in the presence of 25 mM HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> and 5% CO2 on the serosal side. Basal Vt was -22.7 ± 5.4 mV, and Rt was 730 ± 128 Omega  · cm2 (n = 7). The mucosal medium initially either acidified (5 of 7 cultures) or alkalinized (2 of 7 cultures). On average, basal JH was -0.15 ± 0.28 µmol · h-1 · cm-2 (not significantly different from zero). When stimulated with both histamine and ATP, six of seven cultures responded with an increased rate of acidification of the mucosal medium and one culture showed an increase in the rate of alkalinization of the mucosal medium. On average, after stimulation JH was 1.03 ± 0.51 µmol · h-1 · cm-2 (n = 7). Despite the variability in the responses to stimulation, mucosal ZnCl2 consistently blocked JH in all tested cultures by -1.2 ± 0.23 µmol · h-1 · cm-2 (n = 5). After treatment with ZnCl2, three of five treated cultures alkalinized and two acidified the mucosal medium (on average, JH = -0.51 ± 0.33 µmol · h-1 · cm-2; n = 5). All average changes in JH measured in the presence of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2 were not significantly different from the equivalent changes measured in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2-free solutions. These data indicate that in the presence of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2 on the serosal side, the pH of the mucosal medium is determined by the net transport of both acid and base across the cultures. Zinc-sensitive H+ secretion is a significant determinant of the mucosal pH in the presence of serosal HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>/CO2.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

We report here that human airway surface epithelium shows secretion of H+ into the airway lumen regulated by mucosal histamine or ATP. The effect of histamine suggests that the stimulation of H+ secretion in vivo involves mucosal mast cells, which we also show to elicit acid secretion. Thus from our data we propose that during airway inflammation, such as in asthma, in CF, or during allergic reactions, H+ secretion by the airways is activated and the ASL acidifies. In addition, ATP is released by cells under various conditions including airway epithelium during mechanical stress (18). Therefore, stimulants that are unrelated to the mast cell-mediated responses can also lead to acidification of the ASL.

Properties of channel-mediated H+ secretion in airways. We aimed to identify the type of H+ transporter that operates in the apical cell membrane of airways. We used drug sensitivity and voltage- and ion dependence to characterize the H+ transporter. Our data show that H+ secretion by airways occurred primarily via a zinc-sensitive apical membrane H+ conductance for the following reasons. 1) Under ion-free conditions or in the presence of ions rates of H+ secretion were similar. Therefore, the ion-dependent H+ transporters (i.e., the Na+/H+ exchanger and the K+-H+-ATPase) did not contribute significantly. 2) In transepithelial current-clamp experiments the application of positive (outward) current across the epithelium (which polarizes the cytoplasmic face of the apical membrane positively) was accompanied by an increase in H+ secretion. Current passed in the opposite direction reduced H+ currents, indicating that H+ secretion is electrogenic. 3) In whole cell recordings typical identifiers of H+ currents were found including strong outward rectification and activation by depolarization (3, 9, 25, 35). 4) ZnCl2 effectively blocked H+ secretion in transepithelial recordings and in whole cell patch-clamp recordings. In transepithelial recordings Sch-28080, bafilomycin A1, or ouabain had no significant effects on H+ secretion, indicating that the K+-H+-ATPase and the V-type H+-ATPase did not significantly contribute to transepithelial H+ secretion. Thus the results of the blocker experiments are consistent with the results obtained from the ion-free experiments. An apically localized H+ conductance in human airways was identified by its pharmacological and biophysical characteristics, and other transporters could be excluded by the observed ion independence of H+ secretion and by using blockers. Interestingly, in parallel experiments with bovine tracheal cultures we found a significant block of H+ secretion by bafilomycin A1 (data not shown), indicating species-specific variations in H+ transporters in the airways.

Owing to the strong outward rectification of the H+ currents and the localization of the conductance to the apical membrane, this pathway is expected to support H+ fluxes in the secretory direction. This is consistent with our observation that only H+ secretion (and not H+ absorption) was measured in this report (see, for example, Fig. 3). Our data are consistent with the notion that an outward electrochemical H+ gradient exists across the apical membrane that drives H+ secretion across the airway epithelium. H+ conductances in other cell types have been shown to be extremely dependent on the membrane potential such that at negative potentials currents were very small but at depolarizing and positive potentials H+ currents activated. An additional critical feature is the activation of the H+ conductance by an inside-to-outside H+ gradient, as shown in detail for the H+ conductance in rat alveolar cells (8). In fact, in the absence of an H+ gradient the threshold voltage for activation was shown to be nonphysiologically high, current activation was slow, and the currents were very small (8). Thus, in cell types where H+ channels were found, their assumed function is the dissipation of high intracellular H+ concentrations during metabolic acidosis. For example, in phagocytic neutrophils the intracellular space near the membrane acidifies by release of H+ from NADPH during generation of superoxide anions by the membrane-bound NADPH oxidase, and H+ currents are driven by the metabolic acidosis at the membrane (15). Similarly, a H+ conductance in mast cells has been proposed to dissipate the stimulus-induced cytosolic acidification (25). However, in cultured human nasal airway cells cytosolic pH was reported to be 7.15 (30), which under physiological conditions would result in a H+ gradient from the ASL (pH = 6.9) into the cell. Because our data are consistent with an outward H+ gradient, we suggest that the intracellular pH near the apical membrane is markedly acidic in airway cells. This hypothesis relies on the following observations. Mucosal pH of airway cultures equilibrated at pH = 6.85 (Fig. 1). Furthermore, the apical membrane potential (Va) in human airway cell cultures has been reported as -26 ± 1 mV (38) and -19 ± 2 mV (37). Assuming an average Va of -22.5 mV, then at equilibrium the sum of the Nernst potential for H+ and -Va is zero. With a mucosal pH of 6.85 at equilibrium, the intracellular pH is calculated to be 6.5. Currently, however, there is no described source of protons near the apical membrane in airways that would, like the NADPH oxidase in neutrophils, generate acid equivalents.

Proton permeability in airways. Our transepithelial measurements were done at a mucosal pH of 7.3, and peak responses with single agonists were JH = 0.41 µmol · h-1 · cm-2 (Fig. 1). Assuming an intracellular pH of 6.5, Va = -22.5 mV and an amplification of the apical membrane area by the cilia by a factor of 35 (calculated from morphometric data in Ref. 32), then the H+ permeability (P) of the apical membrane during the peak responses is
P=<FR><NU>J<IT> · </IT>(e<SUP><IT>&phgr;</IT></SUP><IT>−</IT>1)</NU><DE>([H<SUP><IT>+</IT></SUP>]<SUB>intra</SUB><IT>−</IT>[H<SUP><IT>+</IT></SUP>]<SUB>muc</SUB><IT> · </IT>e<SUP><IT>&phgr;</IT></SUP>)<IT> · &phgr;</IT></DE></FR>
with phi  = Va · F · R-1 · T-1, where F, R, and T have their usual meanings and [H+]intra and [H+]muc are intracellular and mucosal H+ concentrations, respectively, resulting in P = 9.6 × 10-3 cm/s. In addition, our patch-clamp measurements allowed us to determine the H+ permeability independent of the transepithelial ion fluxes and without the assumptions used above. With the reversal potential of the whole cell currents (-72 mV) and the extracellular pH of 7.3, the apparent intracellular pH is 6.12 in the whole cell experiments. With the specific H+ conductance in JME cells of Gm = 9.2 pS/pF and a specific membrane capacitance of 1 µF/cm2, the H+ permeability for the whole cell patch-clamp experiments is
P=G<SUB>m</SUB><IT> · R · T/</IT>(<IT>F</IT><SUP>2</SUP><IT> · </IT>[H<SUP>+</SUP>]<SUB>m</SUB>)
where [H+]m is the average [H+] in the membrane according to [H+]m = ([H+]muc - [H+]intra)/(ln[H+]muc - ln[H+]intra). This calculation results in a H+ permeability for JME airway cells in whole cell patch-clamp measurements of P = 9.2 × 10-3 cm/s, which is a value closely agreeing with the H+ permeability determined in the transepithelial measurements on primary airway cultures (see above). For comparison, the H+ permeability of other cell types is 2.2 × 10-3 cm/s in Madin-Darby canine kidney membrane vesicles (16), 5 × 10-3 cm/s in renal cortex brush border vesicles (21), and 14 × 10-3 cm/s in alveolar type II cells (8). These calculations indicate that the H+ permeability of airways can be high, and, dependent on the electrochemical driving forces, will permit large H+ fluxes.

Effects of amiloride on apical Na+ channels vs. Na+/H+ exchanger. Amiloride was used to test for the presence of an apically localized Na+/H+ exchanger. Previously, Paradiso (30) found the Na+/H+ exchanger exclusively in the basolateral membrane. We found that H+ secretion was transiently blocked by amiloride in tissues that also expressed a significant amiloride-sensitive Vt. Variable amiloride-sensitive Na+ currents can be expressed by different cultures, depending in part on the culture conditions, source of cells, and experimental conditions (6, 24, 34). Na+ current across the apical membrane is a significant determinant of Va. Thus the measured inhibition of H+ secretion by amiloride is well explained by an amiloride-induced hyperpolarization of Va.

Role of acidic ASL in asthma. In asthma, the pH of condensed exhaled breath has been reported to be markedly acidic (19), a result that could reflect increased rates of acid secretion by the airway epithelium. Asthma is characterized by an increased number of luminal mast cells (12) that release their contents more readily than normal, and the levels of mast cell degranulation products in the ASL are correspondingly higher than normal (10). Thus we propose that increased levels of histamine (and possibly other mast cell products) in the lumen of the airways of asthmatic patients causes an acidification of the ASL. At the histamine-stimulated rates of H+ secretion that we have found (0.41 µmol · h-1 · cm-2) and an estimated buffer capacity of the ASL of 40 mM/pH, the initial rate of acidification of the ASL can be calculated as 0.17 pH units/min. Acidic ASL then produces as yet undetermined functional changes in the airway epithelium that might initiate or exacerbate asthma attacks. Mast cells 1) secrete various factors in addition to histamine (11) and 2) express H+ channels and release H+ (25). Therefore, we tested directly the effect of applying a suspension of the mast cell line HMC-1 to the mucosal surface of tracheal culture. The resulting H+ secretion by the epithelium was significantly larger than that seen with histamine or ATP. However, mast cells alone showed a brief but significant release of acid (Fig. 5), suggesting that the continuously elevated H+ secretion by the airway cultures was mediated by mast cell-derived factors.

Role of acid secretion in CF and bacterial colonization. Asthma and CF share some common symptoms, such as sinusitis and frequent respiratory infections. CF is caused by a mutation in the CFTR Cl- channel. CFTR is also conductive for HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> (31). In addition, Na+ absorption is increased in CF, resulting in a depolarization of Va (37). Both a reduced HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> secretion and a depolarized Va (which would stimulate epithelial H+ secretion) will decrease the pH of the ASL in CF. Preliminary measurements of the mucosal pH in noses of CF patients indeed showed more acidic values than normal (6.2 vs. 6.65; Ref. 7). Currently it is unclear whether the ASL acidity contributes to CF lung disease. However, it has been demonstrated that mucosal acidity affects the function of the epithelium. For example, prolonged treatment of airway epithelium with pH <6.5 caused significant damage (13, 17). Compromised epithelial integrity, in turn, has been shown to cause increased bacterial binding (27). Thus it is possible that ASL acidification is a significant step during airway inflammation and infection in CF as well as asthma.


    ACKNOWLEDGEMENTS

We thank Dr. Walter Finkbeiner (University of California, Davis) for providing the airway cultures, Dr. Terry Machen (University of California, Berkeley) for discussion, and Eric Wunderlich for the JME cell cultures.


    FOOTNOTES

This work was supported by National Heart, Lung, and Blood Institute Grant 1-P50-HL-60288.

Address for reprint requests and other correspondence: H. Fischer, Children's Hospital Oakland Research Institute, 5700 Martin Luther King Jr. Way, Oakland, CA 94609-1673 (E-mail: hfischer{at}chori.org; http://www.chori.org/scientists/fischer.html).

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.

10.1152/ajpcell.00369.2001

Received 2 August 2001; accepted in final form 8 November 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Aris, R, Christian D, Sheppard D, and Balmes JR. Acid fog-induced bronchoconstriction. The role of hydroxymethanesulfonic acid. Am Rev Respir Dis 141: 546-551, 1990[ISI][Medline].

2.   Butterfield, JH, Weiler D, Dewald G, and Gleich GJ. Establishment of an immature mast cell line from a patient with mast cell leukemia. Leuk Res 12: 345-355, 1988[ISI][Medline].

3.   Cherny, VV, and DeCoursey TE. pH-dependent inhibition of voltage-gated H+ currents in rat alveolar epithelial cells by Zn2+ and other divalent cations. J Gen Physiol 114: 819-838, 1999[Abstract/Free Full Text].

4.   Clary-Meinesz, C, Mouroux J, Cosson J, Huitorel P, and Blaive B. Influence of external pH on ciliary beat frequency in human bronchi and bronchioles. Eur Respir J 11: 330-333, 1998[Abstract/Free Full Text].

5.   Coakley, RD, Grubb BR, Gatzy JT, Chadburn JL, and Boucher RC. Differential airway surface liquid (ASL) pH, HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> and K+ homeostasis in cultured human and dog bronchial epithelium (Abstract). Pediatr Pulmonol Suppl 20: 194, 2000.

6.   Coleman, DL, Tuet IK, and Widdicombe JH. Electrical properties of dog tracheal epithelial cells grown in monolayer culture. Am J Physiol Cell Physiol 246: C355-C359, 1984[Abstract].

7.   Davies, MG, Davies JC, Geddes DM, and Alton EW. Nasal epithelial pH is lower in patients with cystic fibrosis (Abstract). Pediatr Pulmonol Suppl 20: 194, 2000.

8.   DeCoursey, TE, and Cherny VV. Deuterium isotope effects on permeation and gating of proton channels in rat alveolar epithelium. J Gen Physiol 109: 415-434, 1997[Abstract/Free Full Text].

9.   Demaurex, N, Grinstein S, Jaconi M, Schlegel W, Lew DP, and Krause KH. Proton currents in human granulocytes: regulation by membrane potential and intracellular pH. J Physiol (Lond) 466: 329-344, 1993[Abstract].

10.   Ennis, M, Turner G, Schock BC, Stevenson EC, Brown V, Fitch PS, Heaney LG, Taylor R, and Shields MD. Inflammatory mediators in bronchoalveolar lavage samples from children with and without asthma. Clin Exp Allergy 29: 362-366, 1999[ISI][Medline].

11.   Galli, SJ, and Costa JJ. Mast cells. In: The Lung (2nd ed.), edited by Crystal RG, and West JB.. Philadelphia: Lippincott-Raven, 1997, p. 929-946.

12.   Gibson, PG, Saltos N, and Borgas T. Airway mast cells and eosinophils correlate with clinical severity and airway hyperresponsiveness in corticosteroid-treated asthma. J Allergy Clin Immunol 105: 752-759, 2000[ISI][Medline].

13.   Giddens, WE, and Fairchild GA. Effects of sulfur dioxide on the nasal mucosa of mice. Arch Environ Health 25: 166-173, 1972[ISI][Medline].

14.   Guba, M, Kuhn M, Forssmann WG, Classen M, Gregor M, and Seidler U. Guanylin strongly stimulates rat duodenal HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> secretion: proposed mechanism and comparison with other secretagogues. Gastroenterology 111: 1558-1568, 1996[ISI][Medline].

15.   Henderson, LM, and Meech RW. Evidence that the product of the human X-linked CGD gene, gp91phox, is a voltage-gated H+ pathway. J Gen Physiol 114: 771-786, 1999[Abstract/Free Full Text].

16.   Hill, WG, and Zeidel ML. Reconstituting the barrier properties of a water-tight epithelial membrane by design of leaflet-specific liposomes. J Biol Chem 275: 30176-30185, 2000[Abstract/Free Full Text].

17.   Holma, B, Lindegren M, and Anderson J. pH effect on ciliomotility and morphology of respiratory mucosa. Arch Environ Health 32: 216-226, 1977[ISI][Medline].

18.   Homolya, L, Steinberg TH, and Boucher RC. Cell to cell communication in response to mechanical stress via bilateral release of ATP and UTP in polarized epithelia. J Cell Biol 150: 1349-1360, 2000[Abstract/Free Full Text].

19.   Hunt, JF, Fang K, Malik R, Snyder A, Malhotra N, Platts-Mills TAE, and Gaston B. Endogenous airway acidification. Implications for asthma pathophysiology. Am J Respir Crit Care Med 161: 694-699, 2000[Abstract/Free Full Text].

20.   Illek, B, and 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].

21.   Ives, HE, and Verkman AS. Effects of membrane fluidizing agents on renal brush border proton permeability. Am J Physiol Renal Fluid Electrolyte Physiol 249: F933-F940, 1985[ISI][Medline].

22.   Jayaraman, S, Song Y, Vetrivel L, Shankar L, and Verkman AS. Noninvasive in vivo fluorescence measurement of airway-surface liquid depth, salt concentration, and pH. J Clin Invest 107: 317-324, 2001[Abstract/Free Full Text].

23.   Jefferson, DM, Valentich JD, Marini FC, Grubman SA, Iannuzzi MC, Dorkin HL, Li M, Klinger KW, and Welsh MJ. Expression of normal and cystic fibrosis phenotypes by continuous airway epithelial cell lines. Am J Physiol Lung Cell Mol Physiol 259: L496-L505, 1990[Abstract/Free Full Text].

24.   Kondo, M, Finkbeiner WE, and Widdicombe JH. Cultures of bovine tracheal epithelium with differentiated ultrastructure and ion transport. In Vitro Cell Dev Biol 29A: 19-24, 1992.

25.   Kuno, M, Kawawaki J, and Nakamura F. A highly temperature-sensitive proton current in mouse bone marrow-derived mast cells. J Gen Physiol 109: 731-740, 1997[Abstract/Free Full Text].

26.   Kyle, H, Ward JP, and Widdicombe JG. Control of pH of airway surface liquid of the ferret trachea in vitro. J Appl Physiol 68: 135-140, 1990[Abstract/Free Full Text].

27.   Lee, A, Chow D, Haus B, Tseng W, Evans D, Fleiszig S, Chandy G, and Machen TE. Airway epithelial tight junctions and binding and cytotoxicity of Pseudomonas aeruginosa. Am J Physiol Lung Cell Mol Physiol 277: L204-L217, 1999[Abstract/Free Full Text].

28.   Lee, MC, Penland CM, Widdicombe JH, and Wine JJ. Evidence that Calu-3 human airway cells secrete bicarbonate. Am J Physiol Lung Cell Mol Physiol 274: L450-L453, 1998[Abstract/Free Full Text].

29.   Luk, CK, and Dulfano MJ. Effect of pH, viscosity and ionic-strength changes on ciliary beating frequency of human bronchial explants. Clin Sci (Lond) 64: 449-451, 1983[ISI][Medline].

30.   Paradiso, AM. ATP-activated basolateral Na+/H+ exchange in human normal and cystic fibrosis airway epithelium. Am J Physiol Lung Cell Mol Physiol 273: L148-L158, 1997[Abstract/Free Full Text].

31.   Poulsen, JH, Fischer H, Illek B, and Machen TE. Bicarbonate conductance and pH regulatory capability of cystic fibrosis transmembrane conductance regulator. Proc Natl Acad Sci USA 91: 5340-5344, 1994[Abstract].

32.   Rhodin, JAG Ultrastructure and function of the human tracheal mucosa. Am Rev Respir Dis 93 Suppl: 1-15, 1966[ISI][Medline].

33.   Smith, JJ, and Welsh MJ. cAMP stimulates bicarbonate secretion across normal, but not cystic fibrosis airway epithelia. J Clin Invest 89: 1148-1153, 1992[ISI][Medline].

34.   Stutts, MJ, Cotton CU, Yankaskas JR, Cheng E, Knowles MR, Gatzy JT, and Boucher RC. Chloride uptake into cultured airway epithelial cells from cystic fibrosis patients and normal individuals. Proc Natl Acad Sci USA 82: 6677-6681, 1985[Abstract].

35.   Thomas, RC, and Meech RW. Hydrogen ion currents and intracellular pH in depolarized voltage-clamped snail neurones. Nature 299: 826-828, 1982[ISI][Medline].

36.   Widdicombe, JG. Relationships among the composition of mucus, epithelial lining liquid, and adhesion of microorganisms. Am J Respir Crit Care Med 151: 2088-2092, 1995[Abstract].

37.   Willumsen, NJ, and Boucher RC. Intracellular pH and its relationship to regulation of ion transport in normal and cystic fibrosis human nasal epithelia. J Physiol (Lond) 455: 247-269, 1992[Abstract].

38.   Willumsen, NJ, Davis CW, and 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].

39.   Wong, CH, Matai R, and Morice AH. Cough induced by low pH. Respir Med 93: 58-61, 1999[ISI][Medline].

40.   Yamaya, M, Finkbeiner WE, Chun SY, and 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].


Am J Physiol Cell Physiol 282(4):C736-C743
0363-6143/02 $5.00 Copyright © 2002 the American Physiological Society