Critical modifier role of membrane-CFTR dependent ceramide signaling in lung injury and emphysema

doi:10.4049/jimmunol.1002850

J Immunol. Author manuscript; available in PMC 2012 January 1.

Published in final edited form as:

J Immunol. 2011 January 1; 186(1): 602–613.

Published online 2010 December 6. doi: 10.4049/jimmunol.1002850

PMCID: PMC3119853

NIHMSID: NIHMS250282

Critical modifier role of membrane-CFTR dependent ceramide signaling in lung injury and emphysema

Manish Bodas,¹ Taehong Min,¹ Steven Mazur,¹ and Neeraj Vij^1,^2,³

¹Department of Pediatric Respiratory Sciences, Johns Hopkins University, Baltimore MD

²Institute of Clinical and Translational Research, Johns Hopkins University, Baltimore MD

³Address for correspondence: Department of Pediatrics, 600 N. Wolfe Street, CMSC 3-122, Baltimore, MD 21287, Email: nvij1/at/jhmi.edu, Phone: 410-955-9783, Fax: 410-502-8007

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Abstract

Ceramide accumulation mediates the pathogenesis of chronic obstructive lung diseases. Although, an association between lack of CFTR and ceramide accumulation has been described, it is unclear how membrane-CFTR may modulate ceramide signaling in lung injury and emphysema. The Cftr^+/+- and Cftr^−/−- mice and cells were used to evaluate the CFTR-dependent ceramide signaling in lung injury. Lung tissue from control and COPD patients was used to verify the role of CFTR-dependent ceramide signaling in pathogenesis of chronic emphysema. Our data reveals a novel finding that CFTR expression inversely correlates with severity of emphysema and ceramide-accumulation in COPD subjects compared to controls. We found that chemical inhibition of de novo- ceramide-synthesis controls Pa-LPS induced lung injury in Cftr^+/+-mice, while its efficacy was significantly lower in Cftr^−/−-mice indicating that membrane-CFTR is required for controlling lipid-raft ceramide levels. Inhibition of membrane-ceramide release showed enhanced protective effect in controlling Pa-LPS induced lung injury in Cftr^−/−- mice as compared to the Cftr^+/+, confirming our observation that CFTR regulates lipid-raft ceramide- levels and signaling. Our results indicate that inhibition of de novo- ceramide-synthesis may be effective in disease states with low-CFTR expression like emphysema and chronic lung injury but not in complete absence of lipid-raft CFTR as in ΔF508-CF. In contrast, inhibiting membrane ceramide release has the potential of a more effective drug candidate for ΔF508-CF but may not be effectual in treating lung injury and emphysema. Our data demonstrates the critical role of membrane-localized CFTR in regulating ceramide-accumulation and inflammatory-signaling in lung injury and emphysema.

Introduction

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 (Pa) bacterial infection has been shown to have critical role in pathogenesis of both CF and COPD (4-6) but it is not clear why these patients are highly sensitive to Pa infections. The absence of cystic fibrosis transmembrane conductance regulator (CFTR) protein from the plasma membrane is known to result in an inherent hyper-inflammatory 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 (42). The earlier studies indicate that genetic mutations in CFTR may be a risk factor for chronic lung diseases like COPD, emphysema and asthma that warrants further investigation (17, 18). In addition, CFTR is known to regulate membrane accumulation of the bioactive lipid, ceramide, that is proposed as a mechanism for pathogenesis of emphysema, COPD (19), CF (7) and chronic lung inflammation (20, 21).

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 H et al shows 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 wt-CFTR controls ceramide accumulation (22). 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 been previously demonstrated that the last 3 amino acids in the COOH-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 (23, 24). 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 (25) and also results in ceramide accumulation.

Our present work supports and expands these important findings and correlates the expression of membrane and lipid-raft (26, 27) localized CFTR to ceramide signaling and severity of lung disease. Our data here shows that CFTR regulates tight junction formation (28), ceramide accumulation and inflammatory signaling in lung injury and emphysema.

Materials and Methods

Reagents and treatments

The cells were cultured at 37°C with 5% CO₂ in MEM [(CFBE4lo-, CFBE4lo-wt-CFTR (from Dr. Dieter Gruenert)], DMEM/F12 (HEK-293) or RPMI-1640 [splenocytes, neutrophils and macrophages] media, supplemented with 10% Fetal Bovine Serum (FBS) and 1% Penicillin, Streptomycin and Amphotericin B (PSA) from Invitrogen. The Pseudomonas aeruginosa LPS (Pa-LPS, Sigma), Fumonisin-B1 (FB1, Cayman Chemicals), Amitriptyline (AMT, Sigma), Methyl-β-cyclodextrin (CD, Sigma), Concanavalin A (ConA, Sigma), TNFα (Invitrogen), and cigarette smoke extract (CSE, Murty Pharmaceuticals Inc) 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 ConA, 10 ng/ml TNFα 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 100μl total volume of PBS, while control mice received PBS alone.

Murine experiments and human subjects

All animal experiments were carried out in accordance with the Johns Hopkins University (JHU) Animal Care and Use Committee (ACUC) approved protocols. We used age, weight and sex matched (24 weeks old), B6- 129S6- Cftr^−/− (Cftrtm^1Kthc–TgN^(FABPCFTR)) (29, 30) and Cftr^+/+ inbred mice strains (procured from Case Western Reserve University Animal Resource Center, 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 is shown. All mice were housed in controlled environment and pathogen-free conditions. We induced lung injury in these mice by i.t instillation of Pa-LPS (20μg in 100 μl PBS) for 12 hours that resulted in approximately 1-2 g loss in body weight. The de novo ceramide synthesis or membrane ceramide release was partially inhibited by i.t (50μg in 100 μl PBS) FB1 or AMT administration, 12 hours post Pa-LPS treatment. Mice were sacrificed 24 hours after drug treatment, and the bronchoalveolar lavage fluid (BALF) was collected for cytokine ELISA’s. The lungs were fixed in 10% buffered formalin phosphate (Fisher Scientific), paraffin embedded and cut into longitudinal sections (5 micron thick) on glass slides for immunostainings. For, depletion of membrane Cftr in murine lungs we used 72-hour i.t. CD treatment, and lung tissues were collected as above. The mice (3-4 mice per group) were exposed to cigarette smoke (CS) using the TE-2 cigarette smoking machine (Teague Enterprises, Davis, CA). The mice (3-4 mice per group, 8 to 10 week old) were exposed to cigarette smoke (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 hours/day for 5 days. An average total particulate matter (TPM) 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 hours after the last CS exposure. The human lung tissue samples from Gold I (mild, FEV1% predicted>80%), Gold II (moderate, FEV1% predicted=50~80%) and Gold Stage III-IV (severe/very severe, FEV1% predicted<50%) non-tumor COPD (with FEV1/FVC ratio of less than 70%) and Gold 0 (at risk) control subjects (procured from LTRC, NHLBI, NIH) were used for quantification and localization of indicated proteins by immunostaining. All the subjects were stable and Gold I-IV subjects had emphysema. Moreover, 1 patient in each group (Gold I-IV) had their first degree blood relatives with chronic bronchitis. A detailed description of the human subjects is shown in Table 1.

Table 1

Patient Characteristics

In vitro and ex vivo experiments

The macrophages and neutrophils from Cftr^+/+ and Cftr^−/− mice were isolated by intraperitoneal (i.p) injection of 1 ml 4% Thioglycollate broth (Fluka). The peritoneal cavity was flushed as indicated after 6 hours (31) (neutrophils) or 4 days (32) (macrophages) with 10 ml of RPMI-1640 media (Gibco) containing 10% FBS (Gibco) and 1% PSA (Gibco) [complete RPMI media]. The lavage was centrifuged at 1200 rpm for 8-10 minutes at 4°C followed by RBC’s lysis in LCK lysis buffer (Quality Biologicals). The 3×10⁵ cells per well were plated in a 6-well plate and cultured overnight in complete RPMI media. The culture supernatants were collected for cytokine ELISA’s and MPO measurements. The spleens were dissected from Cftr^+/+ and Cftr^−/− mice and macerated using the plunger of a 5 ml BD 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 ConA for 72 hours. For splenocyte proliferation assay, 20 μl of the Cell Titer 96® AQ_ueousOne Solution (Promega) was added at 60-hour time point and the plate was incubated at 37°C, 5% CO₂ for another 12 hours. The optical density (OD) at 490 nm was recorded by a 96-well microplate reader (Molecular Devices) using a 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 ConA for 12 hours and the total protein extract was collected using the M-PER protein lysis buffer and 1X protease inhibitor cocktail (Pierce). The human CF bronchial epithelial cells, CFBE41o- and CFBE4lo-wt-CFTR were cultured in MEM media supplemented with 10% FBS (Gibco) and 1% PSA (Gibco). The CFBE4lo-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 number of cells were cultured in glass bottom collagen coated 35 mm petri dishes (Mattek Corporation) and treated for 6 hours with 10 ng/ml Pa-LPS, 5 mM CD and/or 10 ng/ml TNFα. The CFBE4lo-wt-CFTR cells were treated with PBS or 5 mM CD for 24 hours on a 24-well plate and IL-8 secretion in the cell supernatants was quantified by sandwich ELISA (R&D Biosystems). The HEK-293 cells were transiently transfected with wt-CFTR and incubated with increasing doses (0, 40, 80, 120 and 160 μg/ml) of CSE for 12 hours. 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 CFBE4lo-wt-CFTR and CFBE41o- cells treated with PBS, Pa-LPS (10 ng/ml) or FB1 (50μM) for 24 hours. 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) constructs (23) using Lipofecatmine 2000 (Invitrogen) for a total of 48 hours. The cells were treated with 100 μg/ml CSE for the final 12 hours 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 E. coli LPS (Molecular Probes) for the final 3 hours and analyzed by flow cytometry without permeabilizing the cells. The same set of transfections were also performed with or without 50 ng/ml TNFα treatment for 6 hours and lipid-raft proteins were isolated to detect CFTR expression by western blotting.

Immunofluorescence microscopy and flow cytometry

The longitudinal tissue sections from murine or human lungs, or CFBE41o- and CFBE4lo-wt-CFTR cells were immunostained with the primary antibodies (1:50 to 1:200 dilution) for CFTR (rabbit polyclonal, Santa Cruz Biotechnology (scbt), ceramide (mouse monoclonal, Alexis Biochemicals), FoxP3 (rabbit polyclonal, scbt), NFκB (rabbit polyclonal, scbt), Zona occludens -1 (ZO-1) (rabbit polyclonal, scbt), ZO-2 (goat polyclonal, scbt), and neutrophil marker NIMP-R14 (rat monoclonal, Abcam) followed by the secondary antibodies (1:200 dilution), using our previously described protocol (33). The secondary antibodies used were goat anti-rabbit IgG FITC (scbt), 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). Nuclei were detected by Hoechst (Invitrogen) staining while H&E was used to evaluate lung morphology and inflammatory state. Images were captured by Axiovert 200 Carl Zeiss Fluorescence microscope using the Zeiss Axiocam HRC camera and Axiovision software. The membrane localization of ZO-1 and ceramide in CFBE4lo-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 (40X, confocal and 63X, fluorescence) and air (20X and 40X, fluorescence) as the imaging medium. The magnifications for confocal (con) and fluorescence (flr) microscope were EC Plan- Neo Fluar (40X/1.3 oil, con), LD Plan- Achroplan (20X/0.40 Korr Phz, flr), LD Plan- Neo Fluar (40X/0.6X Phz Korr, flr) and LD Plan- Achromat (63X/1.4 oil), respectively with 1.6X optivar. Splenocytes were isolated from Cftr^+/+ and Cftr^−/− mice for flow cytometry and non-specific antibody 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, scbt), and CFTR or intracellular FoxP3 primary antibodies followed by anti-rabbit FITC secondary antibody or stained with CD4-PE followed by intracellular IFNγ-FITC (rat polyclonal, Invitrogen). The macrophages and neutrophils were double stained with the respective cell surface markers, Mac 3 (rat monoclonal, Abcam) or NIMP R-14 (rat monoclonal, Abcam) and ceramide or ZO-1 primary antibodies followed by anti-rat R-PE, anti-mouse Alexa Fluor 488 or anti-rabbit FITC secondary antibodies. The cells were stained and washed 2-times in FACS buffer and resuspended in 0.1 % paraformaldehyde (USB Corporation). Appropriate secondary antibody controls were used in all the flow cytometry experiments. The FIX & PERM®CELL PERMEABILIZATION kit (Invitrogen) was used for IFNγ, FoxP3 and ceramide intracellular staining following the manufacturer’s protocol. The cells were acquired using the BD FACS Caliber instrument and analysis was done by the BD Cell Quest Pro software.

ELISA, MPO activity and reporter assay

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 eBiosciences) following manufacturer’s instructions. MPO levels in neutrophil culture supernatant or mouse BALF were similarly quantified using the MPO ELISA kit (Hycult Biotechnology). For reporter assays, CFBE4lo- wt-CFTR or CFBE4lo- cells were transfected with NFκB firefly luciferase promoter (pGL2) and renila luciferase (pRLTK) control using Lipofectamine 2000 (Invitrogen). Renila luciferase was used as an internal control for normalization of DNA and transfection efficiency of reporter constructs. Cells were induced with 10 ng/ml TNFα and/or 50 μM FB1 for 12 hours and luciferase activities were measured after overnight treatment using the. Dual-Luciferase® Reporter (DLRTM) Assay System (Promega) as described before (27). Data was normalized with internal renila luciferase control for each sample and the changes in reporter activities with CFTR over expression were calculated.

Immunoblotting and lipid-raft isolation

Splenocytes from Cftr^+/+ and Cftr^−/− mice were isolated and stimulated with 5μg/ml ConA for 12 hours. Cells were washed in PBS and total protein was isolated using the 1X M-PER Mammalian protein extraction reagent (Pierce) supplemented with protease inhibitor cocktail (Sigma). The protein lysate was immunoblotted for FoxP3 primary (scbt) or β-actin (Sigma) loading control and anti-rabbit IgG HRP secondary antibodies (Amersham) 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) or β-actin (Sigma) loading control and anti-rabbit or anti-mouse-HRP antibody 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 cocktail. The lung lysate was immunoprecipitated with CFTR 169 antibody (rabbit polyclonal), followed by western blot with CFTR (M3A7) antibody (Abcam). For lipid-raft isolation, CFBE41o- and WT-CFTR CFBE41o cells were plated in 25 cm² tissue culture flask and treated with Pa-LPS (10 ng/ml) and/or FB1 (50 μM) for 24 hours. The cells were washed with cold PBS and raft proteins were isolated using the Signal Protein Isolation kit (G Biosciences). The lung tissue from air and CS exposed mice was similarly harvested in SPE buffer-I and subjected to raft isolation as described below. Briefly, the cells or lung tissue were resuspended in signal protein extraction (SPE) buffer-I and sonicated for 10 seconds to disrupt the cells or tissue. Total protein was quantified in each sample, and equal amount of protein (cells, 300μg and lung tissue, 500μg) 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 FPS (focus protein solubilization) buffer, and used for immunoblotting of ZO-2 (goat primary and anti-goat IgG HRP) and α-actin (rabbit primary and anti-rabbit IgG HRP). The raft protein from mouse lungs or HEK 293 cells was immunoblotted with CFTR 570 antibody (mouse polyclonal antibody, procured from UNC and Cystic Fibrosis Foundation under a MTA).

Statistical analysis

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

Results

CFTR regulates innate- and adaptive- immune response

In order to confirm and expand the hypothesis that functional CFTR is a critical regulator of inflammatory signaling (27), we compared the immune profile of the gut-corrected Cftr^−/− mice to the Cftr^+/+. We quantified the constitutive levels of pro-inflammatory 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^−/− as compared to the Cftr^+/+ (Fig. 1A). We also found a significant increase (p<0.01) in constitutive neutrophil-MPO levels (Fig.1B) in Cftr^−/− as compared to the Cftr^+/+ that 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 bronchoalveolar lavage fluid (BALF) of Cftr^−/− mice as compared to the Cftr^+/+ (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 non-activated splenocytes but Concanavalin A (ConA) induced a significantly higher (**p<0.01, ***p<0.001) splenocyte proliferation and IL-6 secretion in Cftr^−/− as compared to the Cftr^+/+ (Fig. 1D & E). We confirmed that CFTR is expressed on murine splenocytes (Fig 1F, i). The CFTR-deficient splenocytes demonstrate higher numbers of CD4⁺IFNγ⁺ T cells (Fig 1F, ii) supporting the notion that the absence of CFTR results in a constitutive hyper-inflammatory state by inducing the pro-inflammatory response. In addition, prevalence of regulatory T cells is reported in the hyper-inflammatory COPD lungs (34). 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%) as compared to the Cftr^+/+s (0.32%) (Fig 1F, iii). We also confirmed this by FoxP3 immunostaining and western blotting in lung sections and splenocytes, respectively (Fig 1G, H). The data substantiates the previous observations (27, 35-39) and strongly suggests that CFTR is a critical regulator of both innate and adaptive immune responses.

Figure 1

CFTR regulates innate and adaptive immune responses

CFTR expression in inflammatory cells inversely correlates with the levels of ceramide and lipid-raft marker (ZO-1)

Ceramide is a critical regulator of inflammatory and apoptotic signaling (19) and mediates these processes in lung injury (40), asthma (20), emphysema, COPD (19) and CF (7). Moreover, CFTR is present in the lipid-rafts (26, 41), and its role in regulating TNF-R1 and lipid-raft signaling is examined previously (26). 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 shows that macrophages from Cftr^−/− mice have significantly higher ceramide levels as compared to the Cftr^+/+s (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 but 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, 42). Our data indicate a novel 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.

Figure 2

Ceramide and zona occludens 1 (ZO-1) expression is elevated in Cftr^−/− mice immune cells

CFTR regulates membrane ceramide signaling and pathogenesis of chronic emphysema

Ceramide upregulation was recently correlated with emphysema (19) 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 and found that CFTR expression significantly decreases with disease severity while ceramide levels increase (Fig 3A & B, 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. This data implies that lipid-raft localization of CFTR (Fig 3A, insight) controls ceramide accumulation and possibly severity of emphysema. We confirmed our findings in HEK-293 cells transfected with wt-CFTR and show that cigarette smoke extract (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 antibody concentration (Fig 3C, right panel). Extending our findings in the murine model (C57BL/6 mice), we found that acute cigarette smoke exposure (5 hours/day for 5 days) diminished Cftr expression both in the mouse lung lysate (Fig 3D upper panel and Fig 3E left panel, p<0.01) and 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 co-localized 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 (27), the present data verifies that decreased cell surface and lipid-raft expression of CFTR correlates with the increased inflammation and emphysema (Fig 3A, H&E stainings).

Figure 3

Severity of inflammatory lung disease inversely correlates with the membrane-CFTR levels

CFTR expression regulates ceramide signaling in lung injury

In order to verify if 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 (intratracheally, i.t.) for 12 hours, followed by either FB1 or Amitriptyline (AMT) (50 μg/mouse) for another 24 hours. 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, 19). We measured BALF cytokines, IL-6 and IL-1β in all the groups, as a marker of Pa-LPS induced pro-inflammatory insult and the efficacy of the drugs. We found that inhibition of de novo ceramide synthesis by FB1 in Cftr^+/+ mice shows a twofold reduction (p<0.05) in the Pa-LPS induced IL-6 levels (Fig 4A, i), and a very significant decrease (p<0.001) in IL-1β secretion (Fig 4A, ii). In the absence of Cftr (Cftr^−/− mice), FB1 treatment decreased Pa-LPS induced IL-6 (Fig 4A, iii), but the magnitude of rescue was not as efficient as in Cftr^+/+ mice. In addition, IL-1β levels were unaltered by FB1 treatment in the Cftr^−/− mice (Fig 4A, iv). This was also verified by immunostaining of lung sections from these mice for ceramide, NFκB and neutrophil marker, NIMP-R14 (Supplementary Fig 1A, B).

Figure 4

CFTR regulates de novo and membrane ceramide signaling

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 4B, i&ii) while 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 4B, iii&iv). The ceramide, NFκB and NIMP-R14 immunostaining of murine lungs verified these findings (Supplementary Fig 2A, B). Our data concur with findings of Volker et al. (7) who showed that normalization of Asm levels by AMT treatment or partial genetic deficiency reduced pulmonary ceramide levels that protected Cftr-deficient mice from Painfection. 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-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 the potential application as a more effective drug candidate for ΔF508-CF that may not be effectual in treating lung injury and emphysema.

CFTR expression negatively regulates membrane ceramide and lipid-raft signaling

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 CFBE4lo-wt-CFTR and CFBE4lo- cells ± 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 (40, 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 CFBE4lo- cells, we observed higher basal expression of ZO-2 as compared to CFBE4lo-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 this 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 as compared to the Cftr^+/+ mice, which was significantly enhanced by Pa-LPS treatment. Moreover, ceramide was co-localized with ZO-1 indicating its presence in the membrane lipid-rafts. (Fig 5C & D).

Figure 5

CFTR regulates lipid-raft expression and signaling via ceramide

Lack of PDZ-binding domain modulates CFTR-dependent ceramide accumulation

Our data demonstrates the importance of cell surface and lipid-raft CFTR in regulating ceramide mediated inflammatory signaling. The COOH- terminal PDZ-interacting domain of CFTR protein is crucial for its apical membrane polarization and functional robustness (23, 24). In order to investigate the role of this domain in CFTR-dependent inflammatory responses, we over-expressed 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). Since CFTR has been described as a pattern recognition molecule (PRM) for LPS binding (25), our data demonstrates that the PDZ-binding domain of CFTR may be crucial for its function as a PRM. We also demonstrate that expression of ΔTRL-CFTR leads to less CFTR protein reaching the lipid-raft fraction (Figs 6C & D, a & b, a=30 seconds exposure, b=20 min exposure). Treatment with TNFα induces the localization of CFTR to the lipid-rafts but ΔTRL-CFTR mutation compromises its translocation to lipid-raft. Our data suggest PDZ-binding domain is required for CFTR membrane stability and lipid-raft localization. We anticipate binding to PDZ domain containing proteins (ZO-1/2) may be critical for this process.

Figure 6

The PDZ-interacting domain of CFTR regulates ceramide accumulation. (A)

Discussion

We and others have recently shown that apical lipid-raft-localized functional wt-CFTR is critical for controlling the innate immune response (7, 27, 35, 38, 44). Although the link between CFTR dysfunction and inflammatory pathophysiology of CF lung disease remains controversial (45), recent work clarifies and discusses these findings that we have recently reviewed in detail (11, 27). Here, we verify that CFTR is not only critical for regulating the innate immune response in epithelial cells but also regulate adaptive immune response as lack of functional CFTR confers a hyper-inflammatory phenotype to the splenocytes. It has been reported that CD4⁺ T cells from CF patients, have lower IFNγ response (46). We report here that mouse CD4⁺ T cells lacking CFTR (Cftr^−/−) secrete higher amounts of IFNγ as compared to the Cftr^+/+. A recent study by Svetlana O et al showed that although natural T regulatory cells (T regs) were increased in Pa infected Cftr^+/+ mice, depletion of T regs did not alter the disease outcome (47). 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 T regs. The lungs of COPD patients similarly harbor higher number of T regs that are proposed to be involved in controlling pulmonary inflammation or autoimmunity (48). We anticipate that similar mechanism may be triggered in the absence of functional CFTR and strategies directed to modulate functional T regs to revert acute or chronic lung disease warrants further investigation (49, 50).

The pro-inflammatory response in the Cftr-deficient mice is known to be mediated by neutrophils and macrophages, the primary cells of the innate immune response (39, 51-54). We evaluated if the defect in lipid metabolism in the absence of CFTR (7) extends to these immune effector cells. For these studies we used the common P. aeruginosa-LPS (Pa-LPS) induced acute lung injury model (55) that is also a component of air pollutants that cause lung inflammation (56). Interestingly, we observed increased ceramide staining in macrophages (Fig 2A, left panel) but not neutrophils (Fig 2B, left panel) from uninfected Cftr^−/− mice, which correlated with the higher constitutive and Pa-LPS induced pro-inflammatory 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 supports the recent findings that CFTR inhibition by CFTR siRNA in human alveolar macrophages renders them a pro-inflammatory phenotype along with an increase in caveolin-1 expression, as it is related to inflammation and apoptosis in 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 as compared to epithelial or other inflammatory cells (52) may account for lack of ceramide accumulation in the Cftr^−/− as compared to Cftr^+/+. Moreover, a recent study inversely correlates CFTR mediated SCN(-) transport to 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 (Pa) 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 pro-inflammatory and pro-apoptotic 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 supports the critical role of membrane or lipid-raft CFTR in ceramide biogenesis and pathogenesis of lung disease (20, 40). This raises the important question if modulation of CFTR expression in airway diseases can contribute to pathogenesis of chronic lung disease. Interestingly, cigarette smoke extract (CSE) is previously shown to inhibit chloride secretion in human bronchial epithelial cells (61). A more direct correlation between CSE and CFTR expression was established by André M. 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 showing that decrease in CFTR expression correlates with severity [Gold 0 (at risk) vs 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 & E). We also found an increase in co-localization of ceramide and zona occludens (ZO)-1 (Fig 3F) in the murine lungs, post 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 this data that ceramide accumulation and chronic Pa 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 (17, 18, 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. 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 cigarette smoke or lack of detection as they die before severe emphysema is developed or 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. We propose, based on our data that the association of apical and lipid-raft CFTR expression with COPD disease severity, ceramide accumulation and signaling has a novel clinical application as both prognostic marker and therapeutics. Further clinical studies are warranted to confirm the role of CFTR as a modifier or pathogenetic susceptibility factor for COPD, emphysema and asthma.

Since ceramide is an important component of lipid-rafts (43), we hypothesized that disruption of raft CFTR by CD (27) may trigger ceramide accumulation and NFκB activation. We selected CD treatment as a method to selectively deplete CFTR from the lipid-rafts over CFTR siRNA 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 NFκB activity, ceramide levels and IL-8 secretion (Supplementary Fig 3A, B). In vivo depletion of lipid-raft Cftr also showed an increase in ceramide, NFκB and neutrophils (Supplementary Fig 3C) levels and activity, confirming our hypothesis that lipid-raft localized CFTR controls ceramide and NFκB mediated pro-inflammatory signaling. We further verified these results by depleting (CD treatment) (27, 41) or inducing (TNFα) (40, 43) lipid-raft CFTR in CFBE4lo-wt-CFTR cells and observed that lipid-raft CFTR expression controls membrane ceramide accumulation (Supplementary Fig 4). Although we understand that CD and TNFα may modulate NFκB signaling by CFTR independent mechanisms that 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 that is known to induce immune receptor clustering and signaling (68). We anticipate this as a potential mechanism by which CFTR regulates ceramide mediated NFκB signaling.

We and others observed that ceramide mediated lung injury and NFκB signaling is prevented by inhibiting de novo ceramide synthesis (FB1) (19), therefore, we tested its efficacy to suppress TNFα induced NFκB reporter activity in the presence or absence of CFTR. FB1 was able to suppress NFκB reporter activity only in the CFBE4lo-wt-CFTR cells but not in the CFBE4lo- cells (Supplementary 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 acid sphingomyelinase (Asm); hence inhibition of de novo ceramide synthesis is rendered ineffective. The importance of 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 Amitriptyline (AMT) rescued the mice from Pa infection. A clinical trial using AMT in CF patients also demonstrates its safety and efficacy as a potent drug candidate (70). In the present 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 NFκB activityand recruitment of neutrophils in the lungs of Cftr^+/+ mice while its inhibitory effect was significantly lower in Cftr^−/− mice indicating that wt-Cftr depletes NFκB activity by controlling TNF-R1 or sphingomyelin (Fig 7). Moreover, treatment with FB1 may not only prevent the ceramide synthesis, it may 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 as compared to 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 while AMT me be more effective in absence of cell surface CFTR like in case of ΔF508-CF.

Figure 7

Schematic of CFTR mediated ceramide signaling

The previous observations that PDZ-interacting domain in CFTR is required for its apical polarization and Cl⁻ channel function (23, 24) lead us to investigate its role in CFTR-dependent ceramide and lipid-raft signaling. Our data demonstrates that the absence of CFTR PDZ-binding domain (ΔTRL) leads to (a) reduction in membrane CFTR levels (Fig 6C), (b) decrease in binding of E. coli LPS to the plasma membrane (Fig 6B) and (c) increased ceramide accumulation, in both constitutive and CSE induced states (Fig 6A). These findings elucidate 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 present 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 novel biomarker for lung injury and emphysema and demonstrate its potential utility as a prognosticator of aforementioned therapeutic strategy. It remains an open question if the development of potent CFTR corrector- (CF-ΔF508) and potentiator- (COPD and emphysema) drug (currently under Phase II-III clinical trial for CF from Vertex Pharmaceuticals) may serve as an effective therapeutic strategy to overcome the ceramide-induced pathological conditions emerging from decreased membrane or lipid-raft CFTR expression. Since Vertex drugs were identified based on their ability to correct CFTR chloride transport function only (71), we anticipate that the development of the 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.

Supplementary Material

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Text

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Acknowledgement

We are thankful to Drs. Dieter Gruenert and William B. Guggino for providing CFBE41o- and CFBE41o-wt-CFTR cell lines and the pGFP-CFTR-ΔTRL construct, respectively. The human lung tissue samples were provided by Lung Tissue Research Consortium, NIH.

Support: Cystic Fibrosis Foundation (R025-CR07 and VIJ07IO), FAMRI, NASA (NNJ06HI17G) and NIH (CTSA UL RR 025005 and RHL096931) grants to NV.

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