Critical Modifier Role of Membrane-Cystic Fibrosis Transmembrane Conductance Regulator-Dependent Ceramide Signaling in Lung Injury and Emphysema Free

Chronic obstructive pulmonary disease (COPD), emphysema, asthma, and cystic fibrosis (CF) subjects suffer from severe tissue-debilitating lung inflammation that is induced by exposure to environmental contaminants like cigarette smoke (CS) and bacterial infections (1–3). The Pseudomonas aeruginosa bacterial infection has been shown to have a critical role in pathogenesis of both CF and COPD (4–6), but it is not clear why these patients are highly sensitive to P. aeruginosa infections. The absence of cystic fibrosis transmembrane conductance regulator (CFTR) protein from the plasma membrane is known to result in an inherent hyperinflammatory lung phenotype causing chronic obstructive lung disease in both human (CF) and Cftr-deficient (Cftr^−/−) mice (7–11). It is evident that CFTR has other critical signaling and/or transport functions (in addition to its well-documented chloride efflux functions) that control the chronic inflammatory response (12–14). In COPD, although inflammatory lung exacerbations cause most of the lung tissue damage, genetic risk factors can modify disease susceptibility. Moreover, emphysema is a disease of the alveoli, and functional CFTR is known to be expressed in alveolar epithelial cells (type I/II) (15, 16) and macrophages (17). The earlier studies indicate that genetic mutations in CFTR may be a risk factor for chronic lung diseases like COPD, emphysema, and asthma, which warrants further investigation (18, 19). In addition, CFTR is known to regulate membrane accumulation of the bioactive lipid ceramide, which is proposed as a mechanism for pathogenesis of emphysema, COPD (20), CF (7), and chronic lung inflammation (21, 22).

Recent data from CF cell lines and Cftr^−/− mice demonstrate that CFTR also acts as a transporter for sphingolipids (13). Moreover, the studies of Hamai (23) show that expression of defective CFTR in lung epithelial cells results in increased mass and synthesis of sphingolipids, including various ceramide species. They demonstrate that expression of wild-type (WT)-CFTR controls ceramide accumulation (23). The authors of these studies propose that deficiency of functional CFTR (Cftr^−/− mice) results in an alteration of the sphingolipid metabolism and an accumulation of cellular ceramide, but how CFTR regulates inflammatory signaling and ceramide accumulation is unclear. It has previously been demonstrated that the last three amino acids in the C terminus of CFTR (T-R-L) comprise a PDZ-interacting domain that is required for the polarization of CFTR to the apical plasma membrane, essential for its chloride channel function (24, 25). We demonstrate here that expression of the mutant form of CFTR lacking the PDZ-interacting domain (ΔTRL), modulates its role as a pattern recognition molecule (26) and results in ceramide accumulation.

Our current work supports and expands these important findings and correlates the expression of membrane and lipid-raft (27, 28) localized CFTR with ceramide signaling and severity of lung disease. Our data show that CFTR regulates tight junction formation (29), ceramide accumulation, and inflammatory signaling in lung injury and emphysema.

The cells were cultured at 37°C with 5% CO₂ in MEM (CFBE41o−, CFBE41o−WT-CFTR [from Dr. Dieter Gruenert, University of California, under material transfer agreement]), DMEM/F12 (HEK-293), or RPMI 1640 (splenocytes, neutrophils, and macrophages) media, supplemented with 10% FBS and 1% penicillin, streptomycin, and amphotericin B (PSA) from Invitrogen (Carlsbad, CA). The P. aeruginosa-LPS (Pa-LPS; Sigma, St. Louis, MO), fumonisin-B1 (FB1; Cayman Chemicals, Ann Arbor, MI), amitriptyline (AMT; Sigma), methyl-β-cyclodextrin (CD; Sigma), Con A (Sigma), TNF-α (Invitrogen), and cigarette smoke extract (CSE; Murty Pharmaceuticals, Lexington, KY) treatments were used for the indicated time points. For in vitro experiments, cells were treated with 10 ng/ml Pa-LPS, 50 μM FB1, 50 μM AMT, 5 mM CD, 5 or 10 μg/ml Con A, 10 ng/ml TNF-α, and/or 0–160 μg/ml CSE as described. Mice were treated by intratracheal (i.t.) instillation with 20 μg Pa-LPS, 50 μg FB1, 50 μg AMT, and/or 50 μg CD as indicated in 100 μl total volume of PBS, and control mice received PBS alone.

All animal experiments were carried out in accordance with Johns Hopkins University (Baltimore, MD) Animal Care and Use Committee-approved protocols. We used age-, weight-, and sex-matched (24 wk old), B6- 129S6- Cftr^−/− (Cftrtm^1Kthc–TgN^(FABPCFTR)) (30, 31) and Cftr^+/+ inbred mice strains (procured from Case Western Reserve University Animal Resource Center, Cleveland, OH; n = 3–5 for all experiments). The changes in cytokine and inflammatory markers between Cftr^+/+ and Cftr^−/− mice were verified by multiple (2, 3) experiments, and representative data are shown. All mice were housed in a controlled environment and pathogen-free conditions. We induced lung injury in these mice by i.t. instillation of Pa-LPS (20 μg in 100 μl PBS) for 12 h, which resulted in ∼1–2 g loss in body weight. The de novo ceramide synthesis or membrane-ceramide release was partially inhibited by i.t. (50 μg in 100 μl PBS) FB1 or AMT administration 12 h after Pa-LPS treatment. Mice were sacrificed 24 h after drug treatment, and the bronchoalveolar lavage fluid (BALF) was collected for cytokine ELISAs. The lungs were fixed in 10% buffered formalin phosphate (Fisher Scientific, Pittsburgh, PA), paraffin embedded, and cut into longitudinal sections (5 μm thick) on glass slides for immunostaining. For depletion of membrane-CFTR in murine lungs, we used 72-h i.t. CD treatment, and lung tissues were collected as above. The mice (three to four mice per group, 8 to 10 wk old) were exposed to CS using the TE-2 cigarette smoking machine (Teague Enterprises, Davis, CA). The CS was generated by burning research-grade cigarettes (3R4F; 0.73 mg nicotine per cigarette) purchased from the Tobacco Research Institute (University of Kentucky, Lexington, KY) for 5 h/d for 5 d. An average total particulate matter of 150 mg/m³ was recorded in real time during the smoking protocols. The control group of mice was exposed to filtered room air, and all the mice were sacrificed 2 h after the last CS exposure. The human lung tissue samples from Gold stage I (mild, forced expiratory volume [FEV1]% predicted >80%), Gold stage II (moderate, FEV1% predicted = ∼50–80%), and Gold stage III–IV (severe/very severe, FEV1% predicted <50%) nontumor COPD (with FEV1/forced vital capacity [FVC] ratio of <70%) and Gold stage 0 (at risk) control subjects (procured from Lung Tissue Research Consortium [Bethesda, MD], National Heart, Lung and Blood Institute, National Institutes of Health) were used for quantification and localization of indicated proteins by immunostaining. All the subjects were stable, and Gold I–IV subjects had emphysema. Moreover, one patient in each group (Gold I–IV) had first-degree blood relatives with chronic bronchitis. A detailed description of the human subjects is shown in Table I.

The macrophages and neutrophils from Cftr^+/+ and Cftr^−/− mice were isolated by i.p. injection of 1 ml 4% thioglycolate broth (Fluka, St. Louis, MO). The peritoneal cavity was flushed as indicated after 6 h (32) (neutrophils) or 4 d (33) (macrophages) with 10 ml RPMI 1640 medium (Life Technologies, Carlsbad, CA) containing 10% FBS (Life Technologies) and 1% PSA (Life Technologies) (complete RPMI medium). The lavage was centrifuged at 1200 rpm for 8–10 min at 4°C followed by RBC lysis in LCK lysis buffer (Quality Biologicals, Gaithersburg, MD). The 3 × 10⁵ cells per well were plated in a 6-well plate and cultured overnight in complete RPMI medium. The culture supernatants were collected for cytokine ELISAs and myeloperoxidase (MPO) measurements. The spleens were dissected from Cftr^+/+ and Cftr^−/− mice and macerated using the plunger of a 5-ml BD (San Diego, CA) syringe. The suspension was subjected to RBC lysis as described above, and 2 × 10⁵ splenocytes per well were cultured in a 96-well plate. The cells were treated with 5 or 10 μg/ml Con A for 72 h. For splenocyte proliferation assay, 20 μl of the Cell Titer 96 AQ_ueousOne Solution (Promega, Madison, WI) was added at the 60-h time point, and the plate was incubated at 37°C, 5% CO₂ for another 12 h. The OD at 490 nm was recorded by a 96-well microplate reader (Molecular Devices, Sunnyvale, CA) using SOFT-MAX-Pro software (Molecular Devices) as a measure of cell proliferation. For immunoblotting, splenocytes (2 × 10⁶ cells/well in a 6-well plate) were treated with 5 μg/ml Con A for 12 h, and the total protein extract was collected using the M-PER protein lysis buffer and 1× protease inhibitor mixture (Pierce, Rockford, IL). The human CF bronchial epithelial cells, CFBE41o− and CFBE41o−WT-CFTR, were cultured in MEM medium supplemented with 10% FBS (Life Technologies) and 1% PSA (Life Technologies). The CFBE41o−WT-CFTR cells were cultured in the presence of 500 μg/ml Hygromycin B (Invitrogen) to maintain the stable expression of WT-CFTR. For fluorescence or confocal microscopy, equal numbers of cells were cultured in glass-bottom, collagen-coated, 35-mm Petri dishes (Mattek, Ashland, MA) and treated for 6 h with 10 ng/ml Pa-LPS, 5 mM CD, and/or 10 ng/ml TNF-α. The CFBE41o−WT-CFTR cells were treated with PBS or 5 mM CD for 24 h on a 24-well plate, and IL-8 secretion in the cell supernatants was quantified by sandwich ELISA (R&D Biosystems, Minneapolis, MN). The HEK-293 cells were transiently transfected with WT-CFTR and incubated with increasing doses (0, 40, 80, 120, and 160 μg/ml) CSE for 12 h. The total protein cell lysate from these samples was extracted as described above, and levels of mature (C form) and immature (B form) CFTR were quantified by Western blotting. The lipid-rafts were isolated from CFBE41o−WT-CFTR and CFBE41o− cells treated with PBS, Pa-LPS (10 ng/ml), or FB1 (50 μM) for 24 h. For the ΔTRL/WT-CFTR experiments, HEK-293 cells were transiently transfected with pEGFP-WT-CFTR or pEGFP-ΔTRL-CFTR (a gift from Dr. William B. Guggino, Johns Hopkins University) constructs (24) using Lipofectamine 2000 (Invitrogen) for a total of 48 h. The cells were treated with 100 μg/ml CSE for the final 12 h and analyzed by flow cytometry. For LPS binding experiment, HEK-293 cells were similarly transfected with WT or ΔTRL constructs and incubated with FITC-labeled Escherichia coli LPS (Molecular Probes, Carlsbad, CA) for the final 3 h and analyzed by flow cytometry without permeabilizing the cells. The same set of transfections was also performed with or without 50 ng/ml TNF-α treatment for 6 h, and lipid-raft proteins were isolated to detect CFTR expression by Western blotting.

The longitudinal tissue sections from murine or human lungs or CFBE41o− and CFBE41o−WT-CFTR cells were immunostained with the primary Abs (1:50 to 1:200 dilution) for CFTR (rabbit polyclonal; Santa Cruz Biotechnology, Santa Cruz, CA), ceramide (mouse monoclonal; Alexis Biochemicals, Plymouth Meeting, PA), Foxp3 (rabbit polyclonal; Santa Cruz Biotechnology), NF-κB (rabbit polyclonal; Santa Cruz Biotechnology), zona occludens (ZO)-1 (rabbit polyclonal; Santa Cruz Biotechnology), ZO-2 (goat polyclonal; Santa Cruz Biotechnology), and neutrophil marker NIMP-R14 (rat monoclonal; Abcam, Cambridge, MA) followed by the secondary Abs (1:200 dilution), using our previously described protocol (34). The secondary Abs used were goat anti-rabbit IgG FITC (Santa Cruz Biotechnology), goat anti-rat IgG (H+L) R-PE, goat anti-mouse IgG/IgM (H+L), Alexa Fluor 488, donkey anti-goat Alexa Fluor 488 (Invitrogen), donkey anti-mouse Dylight 594, donkey anti-rat Dylight 488, and donkey anti-goat Dylight 594 (Jackson ImmunoResearch, West Grove, PA). Nuclei were detected by Hoechst (Invitrogen) staining, and H&E was used to evaluate lung morphology and inflammatory state. Images were captured by an Axiovert 200 Carl Zeiss (Thornwood, NY) Fluorescence microscope using the Zeiss Axiocam HRC camera and Axiovision software. The membrane localization of ZO-1 and ceramide in CFBE41o−WT-CFTR cells was detected by confocal microscopy. The staining protocol for confocal microscopy was similar to the fluorescence staining protocol. The images were captured using a Zeiss LSM 510 Meta confocal microscope and analyzed by Zeiss LSM Image Browser software. All fluorescent and confocal images were captured at room temperature with oil (×40 confocal and ×63 fluorescence) and air (×20 and ×40 fluorescence) as the imaging medium. The magnifications for the confocal and fluorescence microscopes were EC Plan-Neo Fluar (×40/1.3 oil, confocal), LD Plan-Achroplan (×20/0.40 Korr Phz, fluorescence), LD Plan-Neo Fluar (×40/×0.6 Phz Korr, fluorescence), and LD Plan-Achromat (×63/1.4 oil), respectively, with ×1.6 optivar. Splenocytes were isolated from Cftr^+/+ and Cftr^−/− mice for flow cytometry, and nonspecific Ab binding was blocked by incubating them with either donkey or goat serum (1:10; Sigma). Cells were washed once in FACS buffer (2% FBS in PBS) and double stained with CD4-PE (rat monoclonal; Santa Cruz Biotechnology), and CFTR or intracellular Foxp3 primary Abs followed by anti-rabbit FITC secondary Ab or stained with CD4-PE followed by intracellular IFN-γ–FITC (rat polyclonal; Invitrogen). The macrophages and neutrophils were double stained with the respective cell surface markers, Mac 3 (rat monoclonal; Abcam) or NIMP-R14 (rat monoclonal; Abcam) and ceramide or ZO-1 primary Abs followed by anti-rat R-PE, anti-mouse Alexa Fluor 488, or anti-rabbit FITC secondary Abs. The cells were stained and washed two times in FACS buffer and resuspended in 0.1% paraformaldehyde (USB, Cleveland, OH). Appropriate secondary Ab controls were used in all the flow cytometry experiments. The Fix & Permcell Permeabilization kit (Invitrogen) was used for IFN-γ, Foxp3, and ceramide intracellular staining following the manufacturer’s protocol. The cells were acquired using the BD FACSCaliber instrument, and analysis was done with the BD Cell Quest Pro software.

The BALF and cell culture supernatants (n = 3–5) were quantified in triplicate for mouse IL-6, IL-1β, or human IL-8 using ELISA kits (R&D Systems, or eBioscience, San Diego, CA) following the manufacturer’s instructions. MPO levels in neutrophil culture supernatant or mouse BALF were similarly quantified using the MPO ELISA kit (Hycult Biotechnology, Uden, The Netherlands). For reporter assays, CFBE41o−WT-CFTR or CFBE41o− cells were transfected with NF-κB firefly luciferase promoter (pGL2) and renilla luciferase (pRLTK) control using Lipofectamine 2000 (Invitrogen). Renilla luciferase was used as an internal control for normalization of DNA and transfection efficiency of reporter constructs. Cells were induced with 10 ng/ml TNF-α and/or 50 μM FB1 for 12 h, and luciferase activities were measured after overnight treatment using the Dual-Luciferase Reporter Assay System (Promega) as described previously (28). Data were normalized with internal renilla luciferase control for each sample, and the changes in reporter activities with CFTR overexpression were calculated.

Splenocytes from Cftr^+/+ and Cftr^−/− mice were isolated and stimulated with 5 μg/ml Con A for 12 h. Cells were washed in PBS, and total protein was isolated using the 1× M-PER Mammalian protein extraction reagent (Pierce) supplemented with protease inhibitor mixture (Sigma). The protein lysate was immunoblotted for Foxp3 primary (Santa Cruz Biotechnology) or β-actin (Sigma) loading control and anti-rabbit IgG HRP secondary Abs (Amersham, Piscataway, NJ) and developed using the Super Signal West Pico Chemiluminescent Substrate kit (Pierce). Similarly, the total cell lysates from HEK-293 cells transiently transfected with the WT-CFTR and treated with increasing doses of CSE were immunoblotted with CFTR (Cell Signaling Technologies, Danvers, MA) or β-actin (Sigma) loading control and anti-rabbit or anti–mouse-HRP Ab, respectively. The mouse lung tissue from air and CS exposed mice was homogenized in cold tissue lysis buffer (T-PER; Pierce) supplemented with protease inhibitor mixture. The lung lysate was immunoprecipitated with CFTR 169 Ab (rabbit polyclonal), followed by Western blot with CFTR (M3A7) Ab (Abcam). For lipid-raft isolation, CFBE41o− and CFBE41o-WT-CFTR cells were plated in a 25 cm² tissue culture flask and treated with Pa-LPS (10 ng/ml) and/or FB1 (50 μM) for 24 h. The cells were washed with cold PBS, and raft proteins were isolated using the Signal Protein Isolation kit (G Biosciences, Maryland Heights, MO). The lung tissue from air and CS exposed mice was similarly harvested in signal protein extraction (SPE) buffer-I and subjected to raft isolation as described below. Briefly, cells or lung tissue were resuspended in SPE buffer-I and sonicated for 10 s to disrupt the cells or tissue. Total protein was quantified in each sample, and equal amount of protein (cells, 300 μg; and lung tissue, 500 μg) was used to purify the raft fraction. The SPE buffer-II was added followed by incubation on ice for 15 min with intermittent vortexing. The lysate was centrifuged at 20,000 × g for 15 min and the supernatant discarded. The pellet containing signal proteins was solubilized in adequate amount of focus protein solubilization buffer and used for immunoblotting of ZO-2 (Santa Cruz Biotechnology, goat primary and anti-goat IgG HRP) and α-actin (Sigma, rabbit primary and anti-rabbit IgG HRP). The raft protein from mouse lungs or HEK-293 cells was immunoblotted with CFTR 570 Ab (mouse polyclonal Ab; procured from University of North Carolina, Chapel Hill and Cystic Fibrosis Foundation Therapeutics under a material transfer agreement).

Data are represented as the mean ± SEM of at least three experiments, and Student t test and ANOVA were used to determine the statistical significance. The murine and human microscopy data were analyzed by densitometry (MATLAB R2009b; Mathworks, Natick, MA) followed by Spearman’s correlation coefficient analysis to calculate the significance among the indicated groups.

To confirm and expand the hypothesis that functional CFTR is a critical regulator of inflammatory signaling (28), we compared the immune profile of the gut-corrected Cftr^−/− mice with that of the Cftr^+/+ mice. We quantified the constitutive levels of proinflammatory cytokine IL-6 ex vivo in peritoneal macrophages and neutrophils isolated from Cftr^+/+ and Cftr^−/− mice (n = 3) and found significantly (p < 0.001) higher basal IL-6 levels in Cftr^−/− compared with that in the Cftr^+/+ (Fig. 1A). We also found a significant increase (p < 0.01) in constitutive neutrophil-MPO levels (Fig. 1B) in Cftr^−/− compared with those in the Cftr^+/+, which is indicative of the activated state of neutrophils in the absence of CFTR. We confirmed this in vivo using the murine model and observed a significant increase (p < 0.05) in basal and Pa-LPS–induced MPO levels in BALF of Cftr^−/− mice compared with those in the Cftr^+/+ mice (Fig. 1C). To test the outcome of CFTR deficiency on the adaptive immune response, we quantified differences in cell proliferation and IL-6 secretion in splenocytes from Cftr^+/+ and Cftr^−/− mice. We did not find a significant difference in the nonactivated splenocytes, but Con A induced a significantly higher (**p < 0.01, ***p < 0.001) splenocyte proliferation and IL-6 secretion in Cftr^−/− compared with that in Cftr^+/+ (Fig. 1D, 1E). We confirmed that CFTR is expressed on murine splenocytes (Fig. 1Fi). The CFTR-deficient splenocytes demonstrate higher numbers of CD4⁺IFN-γ⁺ T cells (Fig. 1Fii) supporting the notion that the absence of CFTR results in a constitutive hyperinflammatory state by inducing the proinflammatory response. In addition, prevalence of regulatory T cells is reported in the hyperinflammatory COPD lungs (35). We compared the expression of Foxp3 in Cftr^+/+ and Cftr^−/− mice and found constitutively higher numbers of CD4⁺Foxp3⁺ splenocytes in the Cftr^−/− (0.55%) compared with that in the Cftr^+/+ (0.32%) (Fig. 1Fiii). We also confirmed this by Foxp3 immunostaining and Western blotting in lung sections and splenocytes, respectively (Fig. 1G, 1H). The data substantiate the previous observations (28, 36–40) and strongly suggest that CFTR is a critical regulator of both innate and adaptive immune responses.

Ceramide is a critical regulator of inflammatory and apoptotic signaling (20) and mediates these processes in lung injury (41), asthma (21), emphysema, COPD (20), and CF (7). Moreover, CFTR is present in the lipid-rafts (27, 42), and its role in regulating TNF-R1 and lipid-raft signaling has been examined previously (27). We tested the hypothesis that CFTR may be regulating inflammatory signaling via ceramide by inhibiting the formation of membrane and lipid-raft platforms, which would hamper proper clustering of signaling receptor complexes on the plasma membrane. Evidence from previous studies (7) and our data show that macrophages from Cftr^−/− mice have significantly higher ceramide levels compared with those of the Cftr^+/+ mice (Fig. 2A, left panel), which concurs with increased expression of lipid-raft marker ZO-1 (Fig. 2A, right panel). Although the Cftr^−/− neutrophils show a similar increase in ZO-1 expression, ceramide levels remain unchanged (Fig. 2B). We speculate that other mechanisms may be involved in constitutive increase of neutrophil (MPO) activity in the absence of CFTR (14, 17). Our data indicate a mechanism by which CFTR regulates lipid-raft signaling and inflammatory cell function(s). The constitutive defect in the absence of CFTR compromises the ability of these inflammatory cells to respond to infection or injury resulting in pathogenesis of chronic lung disease.

Ceramide upregulation was recently correlated with emphysema (20), and it is known that CFTR deficiency leads to increased ceramide accumulation and lung injury (7). We verified this observation in lung sections from control (Gold 0, at risk) and COPD (Gold I, mild; II, moderate; and III–IV, severe and very severe emphysema) human subjects (Table I) and found that CFTR expression significantly decreases with disease severity while ceramide levels increase (Fig. 3A, 3B, p < 0.001). Although CFTR is not completely absent in severe COPD lungs, its expression is significantly downregulated. We anticipate this as an outcome of lung injury. These data imply that lipid-raft localization of CFTR (Fig. 3A, inset) controls ceramide accumulation and possibly severity of emphysema. We confirmed our findings in HEK-293 cells transfected with WT-CFTR and show that CSE treatment decreased cell surface expression of CFTR (mature, band C) in a dose-dependent manner (Fig. 3C, left panel). The non-transfected HEK-293 cells do not show the CFTR at this Ab concentration (Fig. 3C, right panel). Extending our findings in the murine model (C57BL/6 mice), we found that acute CS exposure (5 h/d for 5 d) diminished CFTR expression both in the mouse lung lysate (Fig. 3D, upper panel, and Fig. 3E, left panel, p < 0.01) and in the purified lipid-raft fraction (Fig. 3D, lower panel, and Fig. 3E, right panel, p < 0.001). Moreover, we also demonstrate that lungs of CS-exposed mice have significantly (p = 0.004, ρ = 0.9316) increased ceramide accumulation that is colocalized with ZO-1 (Fig. 3F), which implies that CS-mediated decrease in CFTR expression results in lipid-raft ceramide accumulation. Therefore, in accord with our previous observation (28), the current data verify that decreased cell surface and lipid-raft expression of CFTR correlates with the increased inflammation and emphysema (Fig. 3A, H&E staining, bottom panel).

To verify whether CFTR regulates ceramide signaling and outcome of lung injury, we used the Pa-LPS–induced mouse model of lung injury. We treated Cftr^+/+ and Cftr^−/− mice with 20 μg/mouse Pa-LPS i.t. for 12 h, followed by either FB1 or AMT (50 μg/mouse) for another 24 h. We inhibited either the de novo ceramide synthesis (FB1) or membrane-ceramide release (AMT), as they have been shown to mediate the pathogenesis of emphysema and CF lung disease, respectively (7, 20). We measured BALF cytokines IL-6 and IL-1β in all the groups as a marker of Pa-LPS–induced proinflammatory insult and the efficacy of the drugs. We found that inhibition of de novo ceramide synthesis by FB1 in Cftr^+/+ mice shows a 2-fold reduction (p < 0.05) in the Pa-LPS–induced IL-6 levels (Fig. 4Ai) and a very significant decrease (p < 0.001) in IL-1β secretion (Fig. 4Aii). In the absence of Cftr (Cftr^−/− mice), FB1 treatment decreased Pa-LPS–induced IL-6 (Fig. 4Aiii), but the magnitude of rescue was not as efficient as that in Cftr^+/+ mice. In addition, IL-1β levels were unaltered by FB1 treatment in the Cftr^−/− mice (Fig 4Aiv). This was also verified by immunostaining of lung sections from these mice for ceramide, NF-κB, and neutrophil marker NIMP-R14 (Supplemental Fig. 1A, 1B).

In contrast, inhibition of membrane-ceramide release by AMT was unable to rescue Pa-LPS–induced IL-6 or IL-1β secretion in Cftr^+/+ mice (Fig. 4Bi, 4Bii), whereas inhibition of membrane-ceramide in the Cftr^−/− mice showed a significant decrease (p < 0.05) in Pa-LPS–induced IL-6 and IL-1β levels (Fig. 4Biii, 4Biv). The ceramide, NF-κB, and NIMP-R14 immunostaining of murine lungs verified these findings (Supplemental Fig. 2A, 2B). Our data concur with findings of Teichgräber et al. (7) who showed that normalization of acid sphingomyelinase (Asm) levels by AMT treatment or partial genetic deficiency reduced pulmonary ceramide levels that protected Cftr-deficient mice from P. aeruginosa infection. Our results indicate that inhibition of de novo ceramide synthesis (not the release) by FB1 may be effective in disease states with low CFTR expression like emphysema and lung injury but not in total absence of apical or lipid-raft CFTR, for instance in ΔF508-cystic fibrosis (ΔF508-CF), where phenylalanine mutation impairs the folding and trafficking of CFTR to the cell surface. In contrast, inhibition of Asm activity or membrane-ceramide release by AMT has potential application as a more effective drug treatment for ΔF508-CF but may not be effectual in treating lung injury and emphysema.

To elucidate the mechanism by which CFTR regulates lipid-raft signaling, we quantified the expression of tight junction protein ZO-2 in purified raft-protein extract from CFBE41o−WT-CFTR and CFBE41o− cells with or without Pa-LPS or FB1. We found that ZO-2 expression was downregulated by Pa-LPS or FB1, only in the presence of WT-CFTR (Fig. 5A). It is possible that Pa-LPS may induce more recruitment of WT-CFTR to the raft (41, 43), which in turn inhibits raft formation (low ZO-2). FB1 is also able to modulate ZO-2 expression by an unknown mechanism that needs further investigation. Moreover, in the absence of functional CFTR in CFBE41o− cells, we observed higher basal expression of ZO-2 compared with that in CFBE41o−WT-CFTR cells. We also observed that neither Pa-LPS nor FB1 is able to modulate ZO-2 expression in these cells (Fig. 5A). To confirm these data, we stained the lung sections from Cftr^+/+ and Cftr^−/− mice with ZO-2 and found a constitutively higher ZO-2 expression in the Cftr-deficient mouse lungs (Fig. 5B). We also tested another marker of tight junctions, ZO-1, and analyzed it by co-immunostaining with ceramide using the lung sections from Cftr^+/+ and Cftr^−/− mice that were treated with Pa-LPS or PBS. We found a constitutive increase in ceramide levels in the Cftr^−/− mice lungs compared with that in the Cftr^+/+ mice, which was significantly enhanced by Pa-LPS treatment. Moreover, ceramide was colocalized with ZO-1 indicating its presence in the membrane lipid-rafts. (Fig. 5C, 5D).

Our data demonstrate the importance of cell surface and lipid-raft CFTR in regulating ceramide-mediated inflammatory signaling. The C-terminal PDZ-interacting domain of CFTR protein is crucial for its apical membrane polarization and functional robustness (24, 25). To investigate the role of this domain in CFTR-dependent inflammatory responses, we overexpressed WT- or ΔTRL- (CFTR lacking the PDZ binding domain) CFTR-GFP in HEK-293 cells and quantified ceramide levels by flow cytometry. We found that expression of ΔTRL-CFTR triggers higher ceramide accumulation (Fig. 6A, upper panel), which is more prominent upon CSE treatment (Fig. 6A, lower panel). Expression of ΔTRL-CFTR also decreases the binding of E. coli LPS-Alexa Fluor 488 to the plasma membrane (Fig. 6B). Because CFTR has been described as a pattern recognition molecule for LPS binding (26), our data demonstrate that the PDZ binding domain of CFTR may be crucial for its function as a pattern recognition molecule. We also demonstrate that expression of ΔTRL-CFTR leads to less CFTR protein reaching the lipid-raft fraction (Fig. 6Ca, 6Cb [6Ca, 30-s exposure; 6Cb, 20-min exposure], 6D). Treatment with TNF-α induces the localization of CFTR to the lipid-rafts, but ΔTRL-CFTR mutation compromises its translocation to lipid-raft. Our data suggest that PDZ binding domain is required for CFTR membrane stability, and lipid-raft-localization and signaling (Fig. 7). We anticipate binding to PDZ domain-containing proteins (ZO-1/2) may be critical for this process.

We and others have recently shown that apical lipid-raft–localized functional WT-CFTR is critical for controlling the innate immune response (7, 28, 36, 39, 44). Although the link between CFTR dysfunction and inflammatory pathophysiology of CF lung disease has been a subject of debate (45), recent work clarifies and discusses these findings that we have recently reviewed in detail (11, 28). In this study, we verify that CFTR is not only critical for regulating the innate immune response in epithelial cells but also regulates the adaptive immune response as lack of functional CFTR confers a hyperinflammatory phenotype to the splenocytes. It has been reported that CD4⁺ T cells from CF patients have lower IFN-γ response (46). We report in this study that mouse CD4⁺ T cells lacking CFTR (Cftr^−/−) secrete higher amounts of IFN-γ compared with that of the Cftr^+/+. A recent study by Carrigan et al. (47) showed that although natural regulatory T cells (Tregs) were increased in P. aeruginosa-infected Cftr^+/+ mice, depletion of Tregs did not alter the disease outcome. Our original finding shows that lack of functional CFTR was able to modulate Foxp3 expression in the lungs and the peripheral tissues indicative of increased number of Tregs. The lungs of COPD patients similarly harbor higher number of Tregs that are proposed to be involved in controlling pulmonary inflammation or autoimmunity (48). We anticipate that a similar mechanism may be triggered in the absence of functional CFTR, and strategies directed to modulate functional Tregs to revert acute or chronic lung disease warrant further investigation (49, 50).

The proinflammatory response in the Cftr-deficient mice is known to be mediated by neutrophils and macrophages, the primary cells of the innate immune response (40, 51–54). We evaluated whether the defect in lipid metabolism in the absence of CFTR (7) extends to these immune effector cells. For these studies, we used the common Pa-LPS–induced acute lung injury model (55) that is also a component of air pollutants that cause lung inflammation (56). Notably, we observed increased ceramide staining in macrophages (Fig. 2A, left panel) but not neutrophils (Fig. 2B, left panel) from uninfected Cftr^−/− mice, which correlates with the higher constitutive and Pa-LPS–induced proinflammatory cytokine levels. We also observed an increase in ZO-1 staining in both macrophages and neutrophils in the absence of CFTR. Some recent studies support our finding and have shown the expression of tight junction proteins like ZO-2 in human macrophages (57, 58). Our data support the recent findings that CFTR inhibition by CFTR small interfering RNA in human alveolar macrophages renders them a proinflammatory phenotype along with an increase in caveolin-1 expression, as it is related to inflammation and apoptosis of macrophages (59). Although constitutive activation of neutrophils in CF is well documented (44, 60), CFTR expression in neutrophils is a subject of debate. Based on current literature, CFTR expression in neutrophils is either very low or absent. It may be possible that lack of CFTR regulates neutrophil function in a ceramide-independent manner. The lower expression of functional CFTR protein on murine and human neutrophils compared to that of epithelial or other inflammatory cells (52) may account for lack of ceramide accumulation in the Cftr^−/− over Cftr^+/+. Moreover, a recent study inversely correlates CFTR-mediated SCN(−) transport with the MPO activity (14). We anticipate this as a potential mechanism of neutrophil activation in the Cftr^−/− mice that mediates the pathogenesis of chronic lung disease in the presence of Pseudomonas aeruginosa infection or lung injury.

It is proposed that changes in sphingosine and sphingosine-1–phosphate uptake in the absence of CFTR may result in membrane-ceramide accumulation (13) that triggers a proinflammatory and proapoptotic response in the respiratory tract. Ceramide forms membrane platforms and alters small lipid-rafts that consist of sphingomyelin and cholesterol. We anticipate that ceramide accumulation in the absence of CFTR might change the function of proteins in the membrane by altering the composition of sphingomyelin–cholesterol-rich lipid-rafts. In favor of this hypothesis, WT-CFTR expression in Cftr^−/− cells controls ceramide accumulation and inflammatory signaling (13). The data also support the critical role of membrane or lipid-raft CFTR in ceramide biogenesis and pathogenesis of lung disease (21, 41). This raises the important question whether modulation of CFTR expression in airway diseases can contribute to pathogenesis of chronic lung disease. Notably, CSE has previously been shown to inhibit chloride secretion in human bronchial epithelial cells (61). A more direct correlation between CSE and CFTR expression was established by Cantin et al. (62), showing that CSE decreased expression of CFTR- gene, protein, and function in Calu-3 cells. Our in vitro studies in HEK-293 cells transfected with WT-CFTR confirm their observation. We document here the first report to our knowledge showing that decrease in CFTR expression correlates with severity (Gold 0 [at risk] versus Gold I [mild], II [moderate], and Gold III–IV [severe and very severe]) of lung emphysema and ceramide accumulation (Fig. 3A). We verified that acute CS exposure of Cftr^+/+ (C57BL/6) mice decreases cell surface and lipid-raft expression of CFTR in murine lungs (Fig. 3D, 3E). We also found an increase in colocalization of ceramide and ZO-1 (Fig. 3F) in the murine lungs after CS exposure. In support of our findings, a recent study (63) demonstrates that CS induces ceramide accumulation in human bronchial epithelial cells. We anticipate based on these data that ceramide accumulation and chronic P. aeruginosa infections in severe COPD patients (5) and CF (5, 64) may be an outcome of decreased CFTR expression.

The previous clinical studies showing the association of CFTR mutations with asthma and COPD (18, 19, 65, 66) were not conclusive due to lack of sufficient controls. Moreover, only few reports have verified emphysema development in CF subjects (67). Our data suggest the critical modifier role of membrane CFTR and ceramide levels in pathogenesis of severe emphysema. An interesting question here is why CF subjects with ΔF508-mutation, resulting in very low membrane-CFTR levels, do not develop severe emphysema? The paucity of emphysema in ΔF508-CF patients may be due to the absence of other contributors like CS or lack of detection as the patients die before severe emphysema develops or is recognized. Nonetheless, our data suggest that pathogenetic changes in membrane and lipid-raft CFTR may have a modifier function in pathogenesis of COPD and emphysema. Based on our data, we propose that the association of apical and lipid-raft CFTR expression with COPD disease severity and ceramide accumulation and signaling has a clinical application as both prognostic marker and therapeutic. Further clinical studies are warranted to confirm the role of CFTR as a modifier or pathogenetic susceptibility factor for COPD, emphysema, and asthma.

Because ceramide is an important component of lipid-rafts (43), we hypothesized that disruption of raft CFTR by CD (28) may trigger ceramide accumulation and NF-κB activation. We selected CD treatment as a method to deplete selectively CFTR from the lipid-rafts over CFTR small interfering RNA or inhibitor as it would result in an overall decrease of CFTR expression and/or function. We found that depletion of raft-CFTR by CD abrogated its regulatory function, marked by a significant increase in NF-κB activity, ceramide levels, and IL-8 secretion (Supplemental Fig. 3A, 3B). In vivo depletion of lipid-raft CFTR also showed an increase in ceramide, NF-κB, and neutrophil (Supplemental Fig. 3C) levels and activity, confirming our hypothesis that lipid-raft–localized CFTR controls ceramide and NF-κB mediated proinflammatory signaling. We further verified these results by depleting (CD treatment) (28, 42) or inducing (TNF-α) (41, 43) lipid-raft CFTR in CFBE41o−WT-CFTR cells and observed that lipid-raft CFTR expression controls membrane-ceramide accumulation (Supplemental Fig. 4), although we understand that CD and TNF-α may modulate NF-κB signaling by CFTR-independent mechanisms, which warrants further investigation and identification of small molecules that can selectively modulate lipid-raft CFTR expression. Nonetheless, our preliminary studies indicate that membrane-localized WT-CFTR inhibits lipid-raft formation as the expression of lipid-raft marker ZO-1/2 was elevated in the absence of CFTR, which is known to induce immune receptor clustering and signaling (68). We anticipate this as a potential mechanism by which CFTR regulates ceramide-mediated NF-κB signaling.

We and others observed that ceramide-mediated lung injury and NF-κB signaling is prevented by inhibiting de novo ceramide synthesis (FB1) (20); therefore, we tested the efficacy of FB1 to suppress TNF-α–induced NF-κB reporter activity in the presence or absence of CFTR. FB1 was able to suppress NF-κB reporter activity only in the CFBE41o−WT-CFTR cells but not in the CFBE41o− cells (Supplemental Fig. 3D). We speculated that WT-CFTR might be regulating membrane-ceramide levels by its interaction with lipid-raft signaling complex (TNF-R1–sphingomyelin) while FB1 suppresses the de novo ceramide hydrolysis. We anticipated that in the CFTR-deficient scenario, membrane-ceramide accumulation is catalyzed by ASM; hence, inhibition of de novo ceramide synthesis is rendered ineffective. The importance of the ASM pathway in several disease models has been comprehensively reviewed (69). Recently, Teichgräber et al. (7) demonstrated that Cftr^−/− mice induce lung ceramide accumulation via Asm, and its inhibition by AMT rescued the mice from P. aeruginosa infection. A clinical trial using AMT in CF patients also demonstrates its safety and efficacy as a potent drug candidate (70). In the current study, we demonstrate that inhibition of de novo (FB1) or membrane-ceramide (AMT) synthesis/release has differential outcomes in controlling the Pa-LPS–induced lung injury in the presence and absence of CFTR. We found that in the presence of WT-CFTR, inhibition of de novo ceramide synthesis by FB1 inhibits Pa-LPS–induced NF-κB activity and recruitment of neutrophils in the lungs of Cftr^+/+ mice while its inhibitory effect was significantly lower in Cftr^−/− mice indicating that WT-CFTR depletes NF-κB activity by controlling TNF-R1 or sphingomyelin (Fig. 7). Moreover, treatment with FB1 may not only prevent the ceramide synthesis but also deplete sphingomyelin levels. This may indirectly modulate the function of ASM that leads to lower ceramide generation and thereby decreased inflammation in Cftr^+/+ mice. In contrast, inhibition of ASM by AMT showed an enhanced protective effect in controlling the Pa-LPS–induced lung injury in Cftr^−/− mice compared with that in the Cftr^+/+ indicating that inhibition of de novo ceramide synthesis by FB1 can be a more potent therapeutic strategy in lung injury, emphysema, and COPD where CFTR raft expression is depleted but not absent, whereas AMT may be more effective in absence of cell surface CFTR, as in the case of ΔF508-CF.

The previous observations that PDZ-interacting domain in CFTR is required for its apical polarization and Cl⁻ channel function (24, 25) led us to investigate its role in CFTR-dependent ceramide and lipid-raft signaling. Our data demonstrate that the absence of CFTR PDZ binding domain (ΔTRL) leads to 1) reduction in membrane CFTR levels (Fig. 6C), 2) decrease in binding of E. coli LPS to the plasma membrane (Fig. 6B), and 3) increased ceramide accumulation in both constitutive and CSE-induced states (Fig. 6A). These findings elucidate a potential mechanism by which CFTR may be sequestered to the lipid-rafts, where it regulates ceramide-mediated inflammatory signaling. We anticipate binding to PDZ domain-containing proteins (ZO-1/2) is required for CFTR membrane stability and lipid-raft translocation. The current study not only describes the critical role of CFTR in pathogenesis of obstructive lung diseases but also demonstrates the scope of an intervention strategy targeting CFTR-dependent lipid-rafts and ceramide for treatment of lung injury and emphysema. In addition, we evaluate CFTR-dependent lipid-rafts as a biomarker for lung injury and emphysema and demonstrate its potential utility as a prognosticator of the aforementioned therapeutic strategy. It remains an open question whether the development of a potent CFTR corrector (CF-ΔF508) and potentiator (COPD and emphysema) drug (currently under phase II–III clinical trial for CF by Vertex Pharmaceuticals, Cambridge, MA) may serve as an effective therapeutic strategy to overcome the ceramide-induced pathological conditions emerging from decreased membrane or lipid-raft CFTR expression. Because the Vertex drugs were identified based on their ability to correct CFTR chloride transport function only (71), we anticipate that the development of selective strategies to modulate CFTR-dependent lipid-rafts and ceramide signaling as proposed in this study will have a more specific therapeutic outcome for treating the chronic stages of lung disease.

We thank Drs. Dieter Gruenert (University of California) and William B. Guggino (Johns Hopkins University) for providing CFBE41o− and CFBE41o−WT-CFTR cell lines and the CFTR-ΔTRL construct, respectively. The human lung tissue samples were provided by the Lung Tissue Research Consortium, National Heart, Lung and Blood Institute, National Institutes of Health.

Disclosures The authors have no financial conflicts of interest.