. 2011 Jul 8;301(4):L557–L567. doi: 10.1152/ajplung.00094.2011

Enhancement of alveolar epithelial sodium channel activity with decreased cystic fibrosis transmembrane conductance regulator expression in mouse lung

Ahmed Lazrak ^1,⁴, Asta Jurkuvenaite ^1,⁴, Lan Chen ^1,⁴, Kim M Keeling ^2,⁵, James F Collawn ^2,^3,^4,⁵, David M Bedwell ^2,^3,^4,⁵, Sadis Matalon ^1,^2,^3,^4,^5,^✉

PMCID: PMC3191758 PMID: 21743028

Abstract

We sought to establish whether the cystic fibrosis transmembrane conductance regulator (CFTR) regulates the activity of amiloride-sensitive sodium channels (ENaC) in alveolar epithelial cells of wild-type, heterozygous (Cftr^+/−), knockout (Cftr^−/−), and ΔF508-expressing mice in situ. RT-PCR studies confirmed the presence of CFTR message in freshly isolated alveolar type II (ATII) cells from wild-type mice. We patched alveolar type I (ATI) and ATII cells in freshly prepared lung slices from these mice and demonstrated the presence of 4-pS ENaC channels with the following basal open probabilities (P_o): wild-type=0.21 ± 0.015: Cftr^+/−=0.4 ± 0.03; ΔF508=0.55 ± 0.01; and Cftr^−/−=and 0.81 ± 0.016 (means ± SE; n ≥ 9). Forskolin (5 μM) or trypsin (2 μM), applied in the pipette solution, increased the P_o and number of channels in ATII cells of wild-type, Cftr^+/−, and ΔF508, but not in Cftr^−/− mice, suggesting that the latter were maximally activated. Western blot analysis showed that lungs of all groups of mice had similar levels of α-ENaC; however, lungs of Cftr^+/− and Cftr^−/− mice had significantly higher levels of an α-ENaC proteolytic fragment (65 kDa) that is associated with active ENaC channels. Our results indicate that ENaC activity is inversely correlated to predicted CFTR levels and that CFTR heterozygous and homozygous mice have higher levels of proteolytically processed ENaC fragments in their lungs. This is the first demonstration of functional ENaC-CFTR interactions in alveolar epithelial cells in situ.

Keywords: lung slices, single channels, forskolin, trypsin, proteolysis

cystic fibrosis (CF) is an autosomal recessive disease caused by mutations in the CFTR gene that is characterized by high-viscosity mucous resulting in airway obstruction, bronchitis, and Pseudomonas aeruginosa infections. Although the precise causes of CF are still debated, epithelial ion transport has been known to be defective and contribute to the pathogenesis of the disease. Knowles et al. (35) reported that patients with CF have higher levels of Na⁺ absorption in their airways. Subsequent studies have documented higher levels of Na⁺ absorption across well-differentiated cultures of airway epithelial cells lacking CFTR (58).

The amiloride-sensitive epithelial Na⁺ channels (ENaC) play an important role in Na⁺ and fluid homeostasis (50, 56). In the short term, ENaC activity is regulated by changes of the open probability (P_o) and the number (N) of channels at the apical plasma membranes. In the long term, ENaC mRNA and protein levels are regulated by a variety of hormones (such as aldosterone), agents that increase intracellular cAMP levels (such as β₂-agonists and adenylate cyclase simulators), reactive oxygen nitrogen species (such as peroxynitrite and superoxide), and mitogen-activated protein kinases (23, 38, 41, 56, 57). Additionally, endogenous channel-activating proteases as well as proteases released by inflammatory cells (trypsin, elastase) activate ENaC either by cleaving critical amino acids in α- and γ-ENaC subunits or by activating signaling pathways (5, 26, 52, 56).

CFTR is a 1,480-amino acid protein and is a member of the traffic ABC family. It functions as a cAMP-regulated Cl⁻ channel (2) and controls other ion conductive pathways including epithelial Cl⁻ and Na⁺ channels (53, 58). Putative mechanisms responsible for the negative regulation of ENaC by CFTR (34) include physical interaction between these two proteins, higher levels of intracellular Cl⁻, and CFTR inhibition of ENaC subunits cleavage by proteases (6, 7, 20, 36).

Recently, Gentzsch et al. (20) showed that CFTR coimmunoprecipitates with ENaC and inhibits its activity by decreasing proteolysis of its extracellular domains. However, these studies were conducted either in Xenopus oocytes with heterologous expression of ENaC and CFTR or in cultured human bronchial cells expressing native ENaC and CFTR. Studies assessing putative interactions between CFTR and ENaC in lung epithelial cells in situ are lacking. These studies are necessary since culture conditions may greatly alter the expression and the activity of various channel proteins that in turn may influence ENaC-CFTR interactions (28, 41).

We utilized novel electrophysiological approaches to establish whether CFTR regulates the activity and proteolysis of ENaC of alveolar epithelial cells in freshly harvested lung slices from wild-type, CFTR heterozygous (Cftr^+/−), knockout (Cftr^−/−), and ΔF508-expressing mice in situ. Furthermore, we assessed the extent of alveolar ENaC activation by forskolin and trypsin in lung slices from these mice. Our results show that ENaC activity inversely correlates to predicted CFTR levels and that CFTR heterozygous and knockout mice have higher levels of proteolytically processed ENaC fragments in their lungs. This is the first demonstration of functional ENaC-CFTR interactions in alveolar epithelial cells in situ.

MATERIALS AND METHODS

Animals.

Mouse strains used were Cftr knockout mice (B6.129P2-Cftr^tm1Unc/J; referred to as Cftr^−/−) (55) and Cftr-ΔF508 mice (B6.129S6-Cftr^tm1Kth/J; referred to as Cftr ΔF508 mice) (64). The Cftr^−/− and ΔF508 mice were backcrossed extensively to C57BL/6 mice to make them congenic on the C57BL/6 background. Heterozygous mice of those lines were bred by personnel of the University of Alabama at Birmingham (UAB) Cystic Fibrosis Center Mouse Genetics Core to produce Cftr^+/+ wild-type, Cftr^+/− heterozygous, and Cftr^−/− or Cftr-ΔF508 homozygous mice used in the present study.

Mice 8 wk of age (∼20 g body wt) were used for experimental protocols. The mice were anesthetized by intraperitoneal injections of diazepam (17.5 mg/kg) and ketamine (450 mg/kg). When totally unresponsive to the pinch test, the thoracic cage was surgically cut open and the lungs were perfused with Krebs-Ringer solution until white in appearance; the lungs were then removed from the thoracic cavity and placed in a beaker containing 20 ml of phosphate-buffered saline (pH 7.4) at 4°C. The UAB Institutional Animal Care and Use Committee (IACUC) approved all experimental procedures involving animals.

Lung slices.

Subsequently, the lungs were washed with Krebs-Ringer solution containing (in mM): 140 NaCl, 3 KCl, 2.5 CaCl₂, 1 MgCl, 10 glucose, and 10 HEPES (pH ≈7.35–7.4). Lung slices were prepared as previously reported (8) with important modifications. In brief, the right lower lobes were dissected and attached to the jig of an OTS 5000 slicer (FHC, Bowdoinham, ME) with cyanoacrylate adhesive gel and sectioned longitudinally with a zirconium blade into 200- to 250-μm-thick slices at 4°C in a temperature-controlled specimen bath (765 HP, FHC). The slices were then transferred to a six-well plate containing ∼2 ml of a DMEM without serum, supplemented with penicillin-streptomycin, and allowed to recover at 37°C in a humidified atmosphere of 5% CO₂-95% air for 3 h.

Electrophysiology.

Lung slices were transferred to a chamber filled with a solution of the following composition (in mM): 130 K-gluconate, 2 MgCl₂, 10 KCl, 10 glucose, and 10 HEPES (pH 7.4, KOH) and held in place with an anchor (Warner Instruments, Hamden, CT). The composition of the pipette solution was as follows (in mM): 140 NaCl, 3 KCl, 2.5 CaCl₂, 1 MgCl, 10 glucose, and 10 HEPES (pH ≈7.35–7.4). For electrophysiology recordings, the chamber was placed on the recording stage of an Olympus microscope EX51WI (Olympus, Center Valley, PA). Single-channel activity in both ATI and type II (ATII) cells was recorded by the cell-attached mode of the patch-clamp technique (21) as previously described (40). The resistance of the pipettes when filled with Krebs solution ranged from 12 to 15 MΩ. Amiloride and forskolin were applied to the cell patch through the pipette, to minimize the damage to the high-resistance seal between the pipette and the cell membrane. In all experiments, the cell membrane patch under the pipette was held at −100 mV using an amplifier (Axopatch 200B, Molecular Devices, Sunnyvale, CA). The holding potential (Vh) across the patch was calculated from the following formula: Vh=Vcell − Vpatch, where Vcell is the cell membrane resting potential and Vpatch is the applied voltage through the pipette. Shlyonsky et al. (54) reported values of −4.4 mV for the holding membrane potentials of ATII cells in situ. Lazrak and Matalon (39) reported values of about −40 mV for the apical membrane differences of human Clara cell-like cells. Since we did not measure the apical membrane potential in our patches, we used a high-potassium solution to depolarize cell membrane potential to near 0 mV. Under these conditions Vh=−Vpatch. Thus, when we applied 100 mV to the pipette, the holding membrane potential across the patch was −100 mV and Na⁺ ions flowed from the pipette into the cell cytoplasm. This approach allows for an accurate calculation of channel conductance. The slices were never reused after exposure to the depolarizing solution. Lung slices were incubated with LysoTracker green (1 μM) for 1 h at room temperature in the experimental chamber to identify ATII cells by the presence of punctate green fluorescence. ATI cell were identified by their characteristic shape.

ENaC Western blotting.

Mice were anesthetized and euthanized. Their lungs were perfused with phosphate-buffered saline until they were clear of blood, removed, and homogenized in radioimmunoprecipitation assay (RIPA) lysis buffer (Thermo Scientific, Rockford, IL) containing protease inhibitors, sonicated for 30 s in ice in 1.5-ml Eppendorf tubes using an ultrasonic liquid processor and centrifuged at 14,000 g for 20 min. Cleared supernatants were used to measure the protein concentration by the BCA assay (Pierce, Rockford, IL) as previously described (14). Equal amounts of protein (150–200 μg) were loaded in 10% Tris·HCl Criterion precast gels (Bio-Rad Laboratories, Hercules, CA); proteins were transferred to polyvinylidene difluoride membranes (Bio-Rad Laboratories) and immunostained with an anti-α-ENaC antibody (Thermo Scientific). Bands were detected by the LumiGLO Western blot kit (Protein Detector, Gaithersburg, MD). To demonstrate antibody specificity, control tissues were incubated with anti-α-ENaC antibodies in the presence of blocking peptides for α-ENaC (Thermo Scientific).

ATII cell isolation.

Mouse ATII cells were isolated as described previously with the following modifications (22, 24). Mice were euthanized and the renal artery and vein were sectioned to drain most of the blood. The chest was opened and the pulmonary vasculature was perfused with normal saline via a catheter inserted into the right ventricle and advanced into the pulmonary artery. The trachea was exposed and cannulated with a trimmed 18-gauge angiocath. A 3-ml syringe containing dispase at 37°C was then attached to the tracheal cannula, and the dispase was pulsed into the alveolar space. The syringe was removed and, after seven drops of the dispase leaked from the cannula into the lungs, 0.45 ml of 45°C low-melting agarose was instilled into the alveolar space and the mouse was packed in ice for 2 min. The lungs were then removed from the chest cavity and incubated in 1 ml of dispase for 45 min at room temperature. The lung tissue was teased away with curved forceps in a culture dish containing DMEM with 0.01% DNase I and then filtered through 100-, 40-, and 25-μm autoclaved nylon mesh. The cells were then spun down and resuspended in DMEM supplemented with 10% fetal bovine serum and plated on cell culture dishes previously coated with biotin-labeled CD45 and CD16 antibodies. The plates were then incubated at 37°C for 2 h. The cell suspensions were poured off the plates, spun down, resuspended again, and counted with a hemocytometer. Cell viability was assessed by Trypan blue exclusion.

RT-PCR.

Total RNA was purified from whole mouse lungs or from highly enriched ATII cells by use of the Ambion RiboPure kit. RNA was DNase treated by use of the Ambion Turbo DNA-free kit. One microgram of RNA was reverse transcribed into cDNA by using oligo(dT) and AMV reverse transcriptase (Promega) in a 50-μl reaction; 5 μl of each RT reaction was used as the DNA template in PCR reactions to amplify either mouse Cftr using primers DB3356 (5′-cag tgg aag agt ttc att ctg-3′) and DB3357 (5′-gca aac ttg gtg atg tcc tg-3′) or mouse Gapdh using primers DB3360 (5′-ttc cag tat gac tcc act cac gg-3′) and DB3361 (5′-tga aga cac cag tag act cca cga c-3′), and 25% of each PCR reaction was visualized on a 1% agarose gel containing Sybr Green.

Statistical analysis.

Data were analyzed via Origin 7.0. Results are reported as group means ± SE. A one-way ANOVA, followed by the Tukey-Kramer test adjusted for multiple comparisons, was used to determine differences among the group means. Differences at P < 0.05 were considered significant.

RESULTS

Lung slices were prepared from five wild-type mice (Cftr^+/+) on 5 separate occasions. Na⁺ single-channel activity was seen in ∼20–25% of patched ATI and ATII cells (5–8 cells per mouse). Mouse ATI and ATII cells expressed two Na⁺ conductances, 4 and 18 pS (Fig. 1). The activity of both channels was totally inhibited by addition of amiloride (10 μM) in the pipette solution (data not shown). Previous reports indicate that the 4-pS channel is highly selective for Na⁺ over K⁺ whereas the 18-pS channel is equal permeable to both cations (28, 41, 63). These findings suggest that both mouse alveolar epithelial cells (ATI and ATII) in situ express similar amiloride-sensitive Na⁺ conductances, as previously reported (27, 32, 43, 45). ATII cells, because of their size and morphology, proved much easier to patch than ATI cells; therefore, we patched ATII cells from wild-type, Cftr^+/−, Cftr^−/−, and ΔF508 littermate mice. We used high-resistance pipettes (15 MΩ) to limit the size of the patch with the intent to minimize the number of channels per patch. Herein, we focused on quantifying the P_o and number of channels per patch of the 4-pS channel in the various groups because of the greater abundance of this conductance relative to the 18-pS conductance.

Fig. 1. — Alveolar type I (ATI) and type II (ATII) cells in wild-type mouse lungs express 2 Na⁺ conductances. A and C: typical records from cell attached patches in ATII and ATI cells, respectively, showing the presence of 2 distinct amplitudes (the first at 0.44 pA and the second at 1.8 pA). The corresponding conductances were 4. 4 and 18 pS. Recordings were obtained at a membrane potential of −100 mV. B and D: corresponding event distribution histograms of the 2 amplitudes. The open probabilities (P_o) of the 2 conductances are shown above their corresponding peaks. These are typical records of data obtained from at least 20 patches from 5 different wild-type mice.

Channel activity at −100 mV and corresponding amplitude histograms of ATII cells for Cftr^+/+, Cftr^+/−, Cftr^−/−, and ΔF508 mice are shown in Figs. 2 and 3 (respectively). Values of P_o for the 4-pS channel varied from 0.21 in Cftr^+/+ mice to 0.8 in Cftr^−/− mice. The data are summarized in Table 1. In some experiments, we incubated slices from Cftr^+/+ mice with the specific CFTR inhibitor N-(2-naphthalenyl)-[(3,5-dibromo-2,4-dihydroxyphenyl)methylene]glycinehydrazide (48) (GlyH-101; 50 μM) for 2 h and then patched ATII cells by using pipettes containing 50 μM GlyH-101. As shown in Table 1, inhibition of CFTR function had no effect on 4-pS P_o, indicating that the presence of CFTR protein is sufficient for inhibition of ENaC activity. Alternatively, residual effects of prior CFTR activity were sufficient to maintain ENaC inhibition under the conditions of this experiment. Finally, as shown in Fig. 4, the P_o of the 18-pS channel in ATII cells of Cftr^−/− mice was more than double its corresponding value in Cftr^+/+. These data indicate that the presence of CFTR affects both the highly Na⁺-selective and cation channels in ATII cells in a similar fashion.

Fig. 2. — Amiloride-sensitive sodium channel (ENaC) activity in ATII cells from *Cftr*^+/+ and *Cftr*^−/+ mice. Lung slices were prepared from the lungs of wild-type and *Cftr*^−/+ mice as detailed in materials and methods; ATII cells were patched in cell-attached mode in situ. A and C: recordings from *Cftr*^+/+ and *Cftr*^−/+ ATII cells, respectively, showing channel openings (downward deflections) for ∼5 min when the membrane patch was held at −100 mV and the cell was depolarized to 0 mV by using high K⁺ in the bathing solution. B and D: corresponding event distribution histograms. The histogram in B (wild-type) shows the activity of 1 channel with amplitude of 0.49 pA, P_o of 0.26, and conductance (g) of 4.9 pS. The corresponding values for the histogram in D (*Cftr*^−/+) were amplitude 0.435; P_o=0.50; g=4.35 pS. Similar recordings were obtained from at least 9 patches from 5 of wild-type and 5 *Cftr*^−/+ mice.

Fig. 3. — ENaC activity in ATII cells from ΔF508 and *Cftr*^−/− mice. Lung slices were prepared from the lungs of as detailed in materials and methods; ATII cells were patched in the cell-attached mode in situ. A and C: recordings showing ENaC activity in ATII cells from ΔF508 and *Cftr*^−/− mice, respectively, for ∼5 min. The membrane patch under the pipette was held at −100 mV and the lung slice was bathed in a high-K⁺ solution. B and D: corresponding all-events distribution histograms. The histogram in B (ΔF508) shows the presence of 1 channel with an amplitude of 0.48 pA, P_o=0.6, and g=4.8 pS. The corresponding values for the histogram in D (*Cftr*^−/−) were amplitude 0.54 pA; P_o=0.86, and g=5.4 pS. Similar recordings were obtained from 5 different mice from each group, with 3–5 recordings per mouse.

Table 1.

Po of the ATII cell 4-pS channel

	Baseline	Forskolin(5 μM)	Trypsin(2 μM)
Cftr^+/+	0.21±0.015	0.7±0.045*	0.66±0.04*
Cftr^+/++GlyH	0.24±0.005
Cftr^−/+	0.39±0.03^†	0.8±0.04*	0.7±0.03*
Cftr^−/−	0.81±0.016^†‡	0.93±0.009*	0.91±0.02*
ΔF508	0.55±0.01^†‡	0.9±0.01*	0.64±0.04

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Values are means ±1 SE; in each case n=9–11 patches from 4 different mice. Po, open probabilities; ATII, alveolar type II cells. +GlyH, ATII cells from Cftr^+/+mice were patched with pipettes containing the cystic fibrosis transmembrane conductance regulator(CFTR) inhibitor GlyH(50 μ). *P < 0.0001 compared with its corresponding baseline(ANOVA followed by Tukey-Kramer test). †P < 0.001 compared with the corresponding Cftr^+/+ value(ANOVA followed by Tukey-Kramer test). ‡P < 0.001 compared with the corresponding Cftr^−/+ value(ANOVA followed by Tukey-Kramer test). Because of the presence of other types of channels in membrane patches, the Po values for the 4-pS channel were computed manually.

Fig. 4. — 18-pS channel activity in ATII cells from *Cftr*^+/+ and *Cftr*^−/− mice. Lung slices were prepared from the lungs of as detailed in materials and methods; ATII cells were patched in the cell-attached mode in situ. A and B: 18-pS channel cation activity in ATII cells from *Cftr*^+/+ and *Cftr*^−/− mice, respectively, for ∼5 min. The membrane patches under the pipette were held at −100 mV and the lung slices were bathed in a high-K⁺ solution. C and D: corresponding all-events distribution histograms from *Cftr*^+/+ and *Cftr*^−/− mice, respectively. The histogram in C shows the presence of 1 channel with an amplitude of 1.8 pA, P_o=0.27, and g=17.7 pS. The corresponding values for the histogram in D (*Cftr*^−/−) are amplitude 1.88 pA; P_o=0.58, and g=18.8 pS. Similar recordings were obtained from 5 different mice from each group, with 3–5 recordings per mouse. E: mean values of P_o (± SE) for *Cftr*^+/+ and *Cftr*^−/− mice; n=4. Values are significantly different from each other (P < 0.05).

In subsequent studies, we added either forskolin (5 μM) or trypsin (2 μM) in the upper half of the pipettes and recorded single-channel activities from ATII cells in situ, prior to and following diffusion of forskolin or trypsin into the cell. As shown in Table 1, both forskolin and trypsin increased the P_o values of the 4-pS channel in ATII cells of wild-type and Cftr^+/− mice. We opted to include forskolin in the solution of the patch pipette since perfusion of lung slices often degraded the integrity of the patches. Significant increases in the number of open channels were also noted in patches of ATII cells from wild-type animals and to a lesser extent from those of ΔF508 (Fig. 5). More importantly, no effect was seen on P_o nor the number of open channels of Cftr^−/− patches, indicating that ENaC was already maximally stimulated (Fig. 5 and Table 1). Western blotting studies also showed higher levels of cleaved 65-kDa ENaC in lung homogenates of Cftr^+/− and Cftr^−/− but not of ΔF508 mice compared with wild-type mice (Fig. 6). On the other hand, levels of the cleaved 37-kDa band in lung homogenates of ΔF508 mice were twice as high as the wild-type mice. All groups had similar levels of total α-ENaC protein (Fig. 6) as well as lung α, β, and γ-Enac mRNA levels (data not shown).

Fig. 5. — Activation of ENaC in ATII cells from *Cftr*^+/+ ΔF508 but not *Cftr*^−/− mice by forskolin. Lung slices were prepared from each group of mice as detailed in materials and methods; ATII cells were patched in the cell-attached mode in situ; pipettes contained the adenylate cyclase activator forskolin (5 μM). A–C: recordings from *Cftr*^+/+, ΔF508, and *Cftr*^−/− mice showing channel openings (downward deflections) for ∼5 min when the cell membrane patch was held at −100 mV and the cell was depolarized to 0 mV by incubating it in a high-K⁺ solution. Notice gradual increase of the current amplitude in A (*Cftr*^+/+) as forskolin diffuses from the pipette in the cell cytoplasm. In contrast, ENaC was more constitutively active in the lungs of *Cftr*^−/− mice and was not activated by forskolin. Solid lines slow the closed state. D–F: corresponding event distribution histograms of 9 open channels for *Cftr*^+/+, 4 for ΔF508 and 1 for *Cftr*^−/−.

Fig. 6. — Lung α-ENaC levels. Equal amounts of lung proteins (150–200 μg) were loaded in 10% Tris·HCl Criterion precast gels, transferred to polyvinylidene difluoride membranes, and immunostained with α-ENaC antibodies (Thermo Scientific, Rockford, IL). Bands were detected by the LumiGLO Western blot kit. A: 1=wild-type; 2=*Cftr*^−/+; 3=*Cftr*^−/−; 4=ΔF508. Representative Western blot was repeated 4 times in lungs from different mice. B: membranes were striped and blotted with equal concentration of an antibody against α-ENaC in the presence of blocking peptide (BP). C: mean values (± 1 SE; n=4) of band density for the indicated molecular masses (in kDa), normalized to α-tubulin. wt, Wild type. D: mean values (1 SE; n=4) of total lung α-ENaC (sum of all bands) normalized to α-tubulin. *P < 0.05 compared with the wild-type value in the same group (2-tailed P test); #P < 0.05 compared with the wild-type value in the same group (1-tailed P test).

We previously reported that only a small fraction of freshly isolated rabbit ATII cells showed evidence of cAMP-activated Cl⁻ channels and none of them showed spontaneous CFTR activity (65). In the present study, cell attached patches of ATII cells with pipettes containing 140 mM NMDG-Cl⁻ failed to show spontaneous channel activity. When forskolin (5 μM) was added to pipettes containing 140 mM NMDGCl⁻, an occasional 6-pS channel was observed consistent with CFTR activity, in addition to larger anion conductances (data not shown). However, RT-PCR on mouse lung homogenates and isolated mouse ATII cells confirmed the presence of Cftr message in wild-type but not in Cftr^−/− mice (Fig. 7).

Fig. 7. — *Cftr* mRNA is expressed in ATII cells. Shown are the products from RT-PCR analysis by using whole wild-type mouse lung or highly purified ATII cells. *Lanes 1*-4 represent reactions using mouse *Cftr* primers. *Lanes 1* and 2 are samples from wild-type^+/+ and *Cftr*^−/− mouse lungs, respectively. *Lane 3* is a sample from highly purified ATII cells isolated from *Cftr*^+/+ mouse lungs. *Lane 4* represents a nontemplate PCR control (ntc). *Lanes 5*-8 represent reactions using mouse *Gapdh* primers from the same samples described in *lanes 1*-4. The *Cftr* PCR product is 158 bp and the *Gapdh* PCR product is 168 bp.

DISCUSSION

Because of the complexity of CFTR-ENaC interactions in native tissues and the difficulties in interpreting the findings, most of the experimental work has been performed in vitro, using heterologous expression systems for the expression of both proteins in either Xenopus oocytes or cell lines. Contradictory results have been reported. For example, Ji et al. (30) reported that Na⁺ currents in Xenopus oocytes injected with α, β, and γ-ENaC were inhibited by coexpression of CFTR and further inhibited when CFTR was activated by cAMP. On the other hand, Lahr et al. (37) reported that the presence of CFTR does not inhibit ENaC in MDCK-C7 cells. Nagel et al. (49) showed that human ENaC is not regulated by CFTR coexpressed in oocytes and concluded that previous reports of CFTR-ENaC inhibition were due to measurement artifacts. Recently, Gentzsch et al. (20) showed that CFTR coimmunoprecipitates with ENaC and inhibits its activity by decreasing proteolysis of its extracellular domains in both Xenopus oocytes and human airway cells. To better understand ENaC-CFTR interactions in their native environment as well as to determine the role of ENaC in the exacerbation of CF pathophysiology in the lungs, we adapted and applied an electrophysiological technique used for studying ion channels in brain slices to mouse lung slices to study ENaC activity in the presence and absence of CFTR.

The present findings are clear evidence for a negative regulatory effect of CFTR on alveolar epithelial ENaC in situ. Furthermore, as shown in Table 1, ENaC P_o values in ATII cells of Cftr^+/− and Cftr^−/− mice were twofold and fourfold higher compared with wild-type controls, respectively. In addition, ENaC P_o values were considerably higher in ΔF508 mice, where CFTR is both defective and not able to reach the plasma membrane. Interestingly, inhibition of CFTR activity did not alter ENaC P_o (or the number of channels present in cell attached patches) in ATII cells of wild-type mice. There are several possible explanations for this result. First, CFTR activity may not be necessary for ENaC inhibition. Second, a small fraction of its basal activity (remaining in the presence of CFTR inhibitor GlyH-101) was sufficient to inhibit ENaC. Finally, CFTR inhibition could not influence the activity of preexisting ENaC that had already been activated.

ATII cells from animals and humans express α-, β-, and γ-ENaC and amiloride-sensitive Na⁺-selective and cation channels (23, 28, 32, 63), and they transport Na⁺ ions in a vectorial fashion (9, 12, 13). To maintain neutrality, Cl⁻ ions must accompany Na⁺ either through transcellular (such as CFTR and other anion channels) or paracellular pathways. However, the existence and function of CFTR in adult ATII cells has been the subject of much debate. ATII cells isolated from rat fetuses and grown to confluence express significant amounts of CFTR on their apical membranes and, when mounted in Ussing chambers, generate basal and cAMP-activated Cl⁻ and HCO₃⁻ currents that are inhibited by glibenclamide but not DIDS (42). It has been argued that CFTR was a remnant from the fetal lung, where it plays an important role in fetal fluid secretion (4).

However, the results of Cheek et al. (11), Nielsen et al. (49a), and more recently Chen et al. (13) and Bove et al. (9) are consistent with the presence of small but detectable basal and cAMP-stimulated anion currents across adult ATII cells that can be inhibited by a variety of CFTR blockers. Furthermore, Chen et al. provided additional evidence for the existence of functional CFTR in rat ATII cells by showing that immunoprecipitated CFTR was in vitro phosphorylated by cAMP and protein kinase A. Recently, Bove et al. demonstrated that cultured human ATII cells immunostained positive with an antibody to CFTR; when mounted in Ussing chambers they exhibited small amounts of cAMP-activated Cl⁻ currents, consistent with the presence of functional CFTR. Shlyonsky et al. (54) reported an increase of whole cell Na⁺ and Cl⁻ currents across ATII cells in lung slices patched in a whole cell mode during cAMP stimulation. On the other hand, we were unable to consistently detect the presence of forskolin-activated Cl⁻ currents in patches of mouse alveolar epithelial cells. We previously reported that only a small fraction of freshly isolated ATII cells showed cAMP-activated Cl⁻ channels when patched in the cell-attached mode (65). Considering the small amount of CFTR present in ATII cells, it will be difficult to identify CFTR single channels in cell-attached patches of these cells.

A functional role for CFTR in alveolar epithelial cells was demonstrated by Fang et al. (17), who showed that intratracheal instillation of β-agonists increased reabsorption of isotonic fluid (an index of vectorial Na⁺ reabsorption) across the distal alveolar spaces of wild-type but not ΔF508 mice. In contrast, our data show that forskolin increased ENaC P_o and the number of channels in ATII cells patches of ΔF508 mice by almost 100%. Other variables such as inflammatory cytokines and reactive oxygen species have been shown to prevent activation of β₂-receptors in vivo (15, 47). Another important difference between our findings and those of Fang et al. is that they reported that ΔF508 had normal levels of alveolar fluid clearance (and thus alveolar sodium transport) whereas our findings indicate significantly higher levels of ENaC activity (the rate-limiting step in this process) in alveolar cells of mice. Thus our findings provide new insight into the regulation of ion transport across distal lung epithelial cells.

The increased Na⁺ absorption through ENaC in CF disease in the proximal airways was proposed more than two decades ago by Knowles et al. (34) when they reported a higher in vivo nasal and tracheobronchial potential difference in CF patients than in normal volunteers. Furthermore, Jiang et al. (31) demonstrated that confluent monolayers of airway cells from patients with CF had higher levels of amiloride-sensitive Na⁺ reabsorption compared with those from normal individuals. Mice overexpressing β-ENaC show symptoms of CF disease without affecting the expression of CFTR (44). Presumably membrane ENaC exists in active and silent pools (38, 56). ENaC activation proceeds by the proteolytic cleavage of both α- and γ-subunits, converting the “silent” pool into active channels (3, 33).

There are at least four proteases suspected to play a major role in the proteolytic activation of ENaC, including furin, which cleaves ENaC before it reaches the plasma membrane (25), and membrane-bound proteases named channel-activating proteases (CAP), which cleave ENaC at the plasma membrane. Vallet et al. (60) cloned a protein-activating protein, CAP1 (prostasin), from A6 cells (Xenopus kidney cells) and demonstrated that it activates ENaC when coexpressed in Xenopus oocytes. Two more proteases, mCAP2 (human TMPRSS4) and mCAP3 (MT-SP1/matriptase) were identified by homology cloning and found to increase the activity of ENaC expressed in oocytes (19). The role of CPA2 in ENaC activation was studied in detail by Garcia-Caballero et al. (19), who showed that this protease cleaves all ENaC subunits at multiple sites including the basic residues close to the degenerin site (close to COOH-terminus of each subunit; α-K561, β-R503, and γ-R515). Our Western blot was performed with an antibody against α-NH₂-terminus. The presence of cleavage sites at the COOH-terminus of α-ENaC explains the presence of the fragments seen in Fig. 6. Trypsin is another protease known to activate ENaC by cleaving specific sites at the α- and γ-subunits (10, 16, 29, 50).

There is little known about the mechanisms involved in the regulation of this process of activation, or when or how the cell determines how many channels are needed at any given moment. In our studies, inclusion of trypsin in the pipette solution increased the activity and P_o of the 4-pS ENaC channels in ATII cells from Cftr^+/+ and Cftr^+/− but not Δ508 or Cftr^−/− mice, suggesting the presence of more silent channels in Cftr^+/+ and Cftr^+/− compared with ΔF508 and Cftr^−/−. Our data suggest that the presence of CFTR may either decrease surface protease activity or obstruct surface proteases' access to critical regions of ENaC subunits. For example, Adebamiro et al. (1) reported that aprotinin inhibited amiloride-sensitive currents across A6 cells. We have previously shown that normal but not oxidized α₁-antitrypsin (α₁AT) decreased ENaC activity in lung epithelial cells and vectorial Na⁺ transport and lung fluid clearance in vivo (40). Reactive species, present in the lungs of mice and humans with CF, are likely to oxidize methionine, a critical residue in the α₁AT active domain, decreasing its antiproteolytic activity. Recently Gentzsch et al. (20) found that CFTR reduced the number of active ENaC channels in Xenopus oocytes and cells, whereas Tarran and colleagues (18, 59) found that airway surface liquid ENaC activity was regulated by proteases. Both findings point to a physiological regulatory mechanisms that maintain ENaC activity under control. However, it is clear that additional mechanisms may account for the increased levels of ENaC activity in the presence of decreased CFTR levels.

We also observed statistically significant differences in ENaC activity between Cftr^+/+ and Cftr^+/− mice (Table 1). Since the heterozygous mice carry a knockout allele on one chromosome, they are expected to have half the CFTR expression of the wild-type mice. Remarkably, the baseline ENaC P_o in the Cftr^+/− mice was 1.9-fold higher than the Cftr^+/+ controls (open probabilities of 0.21 and 0.39, respectively), indicating that ENaC activity was inversely proportional to the level of CFTR expression.

This proportional change in ENaC activity as a function of CFTR expression suggests that CFTR protein inhibits ENaC activation. However, we cannot currently definitely exclude the possibility that a reduction in CFTR activity also contributes to the regulation of ENaC function. Similarly, we found significant differences in ENaC activity measured in lung slices from ΔF508 and Cftr^−/− mice (P_o values of 0.55 and 0.81, respectively). The observed difference between these two CF alleles indicates the remarkable sensitivity of baseline ENaC activity as readout of the CF phenotype. In contrast, forskolin-stimulated ENaC activity was similar between ΔF508 CFTR and Cftr^−/− mice (P_o values of 0.90 and 0.94, respectively). This difference in ENaC activity between the unstimulated and stimulated states shows that the ΔF508 CFTR mice retain a modest ability to regulate ENaC activity, whereas the Cftr^−/− mice do not. It is possible that this regulation is mediated by a small fraction of CFTR-ΔF508 protein that reaches the cell surface. However, this regulation could also occur in the endoplasmic reticulum, where the bulk of CFTR-ΔF508 protein is retained and ultimately degraded. Further studies are required to determine the mechanism by which CFTR regulates ENaC activity.

In humans, higher levels of ENaC lead to CF symptoms, even in the presence of normal CFTR (44). In contrast, ΔF508 and Cftr^−/− mice do not develop CF-related lung disease despite significantly higher ENaC levels. This is because mice either lack goblet cells that secret mucin or compensate for increased Na⁺ absorption across lung epithelial cells by a concomitant increase of Cl⁻ secretion, most likely through Ca²⁺-activated Cl⁻ channels. Unfortunately, we were unable to find convincing electrophysiological evidence for the existence of cAMP-activated Cl⁻ channels in alveolar epithelial cells, a difficulty confirming our previous attempts and studies (65). In the lung, CFTR channels are most likely located in airway cells, which we were unable to patch in our preparations because of the presence of cilia. Our findings indicate that small levels of CFTR expressed in alveolar cells regulate ENaC function.

In the lungs, active Na⁺ transport limits the degree of alveolar edema following damage to the alveolar epithelium; patients with acute lung injury who are still able to concentrate alveolar protein (as a result of active Na⁺ reabsorption) have a better prognosis than those that cannot (46, 61). Intratracheal instillation of phenamil, an irreversible ENaC inhibitor, increases lung water in animals exposed to hyperoxia (62, 63). Conversely, mice with Liddle disease, characterized by ENaC gain-of-function, have higher Na⁺-dependent alveolar fluid clearance and clear hydrostatic edema more effectively than wild-type mice (51). Thus increased vectorial Na⁺ support across lung alveolar epithelia in mice and people with decreased CFTR is both beneficial and detrimental: at the airways, increased fluid reabsorption will increase the viscosity of the mucus leading to obstruction and increased colonization by pathogens; on the other hand, increased alveolar fluid clearance across the distal lungs will help clear alveolar edema and protect from both cardiogenic and permeability-type edema.

GRANTS

This research was supported by National Institutes of Health Grants 5R01HL031197-25, 5U01ES015676-05, 1RO1DK060065-07, and P30DK072482-04.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

ACKNOWLEDGMENTS

The authors thank Laura Cain for technical assistance with the animal experiments and Gloria Son for editorial help with this manuscript.

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PERMALINK

Enhancement of alveolar epithelial sodium channel activity with decreased cystic fibrosis transmembrane conductance regulator expression in mouse lung

Ahmed Lazrak

Asta Jurkuvenaite

Lan Chen

Kim M Keeling

James F Collawn

David M Bedwell

Sadis Matalon

Abstract