We recently demonstrated that the accumulation of ceramide in Cftr-deficient epithelial cells is important for the pathophysiology of CF. However, the role of ceramide in other lung cells, particularly lung macrophages, requires definition. In this study, we report that ceramide is accumulated in Cftr-deficient lung macrophages. Alveolar macrophages contain a vesicle population, which is stained with LysoSensor probes but not by tetramethylrhodamine dextran. These vesicles, presumably secretory lysosomes, exhibit a higher pH in Cftr-deficient macrophages than the corresponding vesicles in lung macrophages isolated from wild-type (WT) mice. Alkalinization of these vesicles in Cftr-deficient macrophages correlates with a failure of the macrophages to respond to infection with various Pseudomonas aeruginosa strains by acutely activating acid sphingomyelinase, releasing ceramide, forming ceramide-enriched membrane platforms that serve to cluster gp91phox, and, most importantly, releasing reactive oxygen species (ROS). In contrast, these events occur rapidly in WT lung macrophages postinfection. Inhibiting ROS in WT macrophages prevents the killing of P. aeruginosa. These findings provide evidence for a novel pH-controlled pathway from acid sphingomyelinase activation via ceramide and clustering of gp91phox to the release of ROS in lung macrophages.

Cystic fibrosis (CF) is the most common autosomal recessive disorder in Western countries, affecting ∼80,000 patients in Europe and the United States. The disease is caused by mutations of the CF transmembrane conductance regulator (CFTR) molecule (1, 2). Genetic defects in the CFTR molecule result in several clinical symptoms, the most common of which are pulmonary problems, especially recurrent and chronic infections with Pseudomonas aeruginosa and Staphylococcus aureus. The reason for the high sensitivity of CF patients to pulmonary infections is unknown.

It has been suggested that CFTR regulates pH in at least some intracellular vesicles (37). The acidification of intracellular vesicles requires high concentrations of protons within the vesicles; these concentrations are achieved by the activity of proton pumps, in particular the v-type adenosine triphosphatase (ATPase) proton pump (8). Because the v-type ATPase protein pump is electrogenic, its activity requires either the accumulation of negatively charged counterions in these vesicles or the active transport of other positively charged ions out of the vesicular lumen (9). Some researchers have proposed that CFTR mediates this influx of counterions into at least some intracellular vesicles (3, 4, 7). However, using internalization-dependent fluorescent dyes or Abs, Verkman and colleagues (10) and Lukacs and associates (11) did not detect any alkalanization of the pH in lysosomes of Cftr-deficient cells.

We have recently shown that an alteration in the pH of at least some vesicles of freshly isolated bronchial epithelial cells lacking Cftr may result in an alteration in sphingolipid metabolism in the respiratory tract of Cftr-deficient mice (5). The alkalinization of these vesicles may result in an imbalance in the activities of the pH-sensitive vesicular enzymes of ceramide metabolism, namely, the activity of acid sphingomyelinase in releasing ceramide from sphingomyelin and the activity of acid ceramidase in degrading ceramide to sphingosine (5). At a pH of 6.0, acid ceramidase is almost inactive, whereas the activity of acid sphingomyelinase is reduced by only 30–40%. This imbalance in activity may result in the chronic accumulation of ceramide in Cftr-deficient cells (5, 12).

In addition, ceramide is acutely released by the acid sphingomyelinase on infection of wild-type (WT) mice or cells with P. aeruginosa infection (1315). In lung alveolar macrophages, P. aeruginosa infection triggers the rapid formation of ceramide-enriched membrane platforms, which cluster and stimulate the activity of NADPH oxidase, thereby resulting in a release of reactive oxygen species (ROS) (14). In this study, we tested the hypothesis that alkalinization of vesicles in Cftr-deficient lung macrophages results in long-term accumulation of ceramide, but prevents the acute activation of acid sphingomyelinase, thereby resulting in defects in the acute response of alveolar macrophages to P. aeruginosa.

Our data identify a distinct population of intracellular vesicles, most likely secretory lysosomes, that exhibit a higher pH in Cftr-deficient macrophages than the corresponding vesicles in WT lung macrophages. Because the signaling pool of the acid sphingomyelinase localizes to secretory lysosomes, we focused in the present manuscript on this LysoSensor Green DND-189 positive, tetramethylrhodamine (TMR)-negative population of acidic vesicles.

Alkalinization of these vesicles correlates with a marked chronic accumulation of ceramide in Cftr-deficient macrophages but not in WT cells. On acute infection with P. aeruginosa, Cftr-deficient macrophages fail to respond adequately by acutely activating acid sphingomyelinase, forming ceramide-enriched membrane platforms, clustering and activating NADPH oxidase within these platforms, releasing ROS, and killing the bacteria, events that are rapidly observed in WT macrophages and that permit WT macrophages to kill P. aeruginosa.

We used the mouse strains B6.129P2(CF/3)-CftrTgH(neoim)Hgu (CftrMHH, syngenic to C57BL/6; kindly provided by Dr. B. Tümmler, Medizinische Hochschule Hannover, Hannover, Germany) and Cftrtm1Unc-Tg(FABPCFTR) (CftrKO; obtained from The Jackson Laboratory, Bar Harbor, ME). Neither of these mouse strains required a special diet. This characteristic is important because some diets have been shown to alter membrane lipid composition (5, 1618). Changes in lipid composition, in particular an alteration in cholesterol and sphingolipids concentrations, may regulate the activity of ion channels and pumps (1921). Littermates of the C57BL/6 mice were used as control animals. All mice were repeatedly tested for the presence of pathogens and were free of any pathogens according to the criteria of the Federation of Laboratory Animal Science Associations.

Lung alveolar macrophages were isolated from WT or CftrMHH mice by bronchoalveolar lavage, as described previously (14). Briefly, the trachea was opened and cannulated with a polyethylene tube. The lung was lavaged with a total of 15 ml ice-cold PBS in 20 aliquots (0.75 ml per aliquot). Approximately 0.5 × 106 to 1 × 106 cells were consistently obtained from each mouse. Cells were pelleted by centrifugation at 300 × g for 15 min, resuspended, and cultured for 1 h in RPMI 1640 (Invitrogen, Karlsruhe, Germany) supplemented with 1 mM HEPES (pH 7.4) in 24-well plates at a density of 105 cells per well. Because alveolar macrophages are extremely adhesive cells, after other blood cells were washed off, we were left with a pure cell culture in which >99% of cells were macrophages, as confirmed by flow cytometry after staining with FITC-coupled anti-CD11b Abs (BD Biosciences, Heidelberg, Germany).

Cellular ceramide levels were measured with a 1,2-diacylglycerol (DAG) kinase assay, as previously described (14). Macrophages were cultured for 60 min and extracted in CHCl3:CH3OH:1 N HCl (100:100:1, v/v/v). Phases were separated, and the lower phase was collected, dried, and subjected to alkaline hydrolysis of DAG in 0.1 N methanolic KOH at 37°C for 60 min. Samples were extracted again, and the lower phase was dried. Samples were resuspended in 20 μl detergent solution consisting of 7.5% (w/v) n-octyl glucopyranoside and 5 mM cardiolipin in 1 mM diethylenetriaminepentaacetic acid (DTPA). Samples were then sonicated for 10 min in a bath sonicator; to these samples was added 70 μl of an assay buffer consisting of 0.1 M imidazole/HCl (pH 6.6), 0.1 M NaCl, 25 mM MgCl2, and 2 mM EGTA; 2.8 mM DTT; 5 μM ATP; 10 μCi of [32P]ATP; 10 μl DAG kinase (diluted in 1 mM DTPA; pH 6.6); and 0.01 M imidazole/HCl. The kinase reaction was performed for 30 min at room temperature, and the samples were extracted in 1 ml CHCl3:CH3OH:1 N HCl (100:100:1, v/v/v), 170 μl buffered saline solution (135 mM NaCl, 1.5 mM CaCl2, 0.5 mM MgCl2, 5.6 mM glucose, 10 mM HEPES; pH 7.2), and 30 μl 100 mM EDTA solution. Phases were separated, and the lower phase was dried. Samples were dissolved in 20 μl CHCl3:CH3OH (1:1, v/v). Lipids were separated on Silica G-60 TLC plates with CHCl3:CH3OH:CH3COOH (65:15:5, v/v/v), and the plates were dried and exposed. Ceramide spots were identified by comigration with a C16-ceramide standard and removed from the plate. The incorporation of [32P] into ceramide was quantified by liquid scintillation counting. Comparison with a standard curve using C16-ceramide permitted the determination of ceramide amounts.

To observe lysosomal alkalinization, we incubated lung macrophages in PBS buffer with or without 100 nM bafilomycin, an H+-ATPase pump inhibitor (Sigma-Aldrich, Deisenhofen, Germany), for 30 min. The cells were then loaded with LysoSensor Green DND-189 (1 μM; Molecular Probes, Invitrogen, Karlsruhe, Germany) in PBS buffer for 15 min at 37°C. Cells were washed twice with PBS and immediately visualized with a Leica SP2 confocal microscope (Leica Microsystems, Wetzlar, Germany). We measured the relative mean fluorescence intensity of at least 200 cells per sample. To obtain a pH standard curve, we loaded cells with LysoSensor Green DND-189 for 15 min at 37°C; the lysosomal pH was equalized by incubating cells for 15 min at 37°C either in PBS containing the ionophore nigericin (10 μM) or in a calibration buffer (120 mM KCl, 20 mM NaCl, 1 mM CaCl2, 1 mM MgCl2, and 10 mM HEPES with the pH adjusted to either 4.5, 5.0, 5.5, 6.0, or 6.5, respectively). We observed no differences in fluorescence between the cells incubated in PBS and those incubated in the HEPES buffers.

To determine whether LysoSensor Green DND-189 and TMR-dextran (10 kDa, Molecular Probes) target the same vesicle population, we incubated lung macrophages with TMR-dextran of various concentrations (1 mg/ml, 200 μg/ml, 50 μg/ml, 20 μg/ml, and 10 μg/ml) for 1 h at 37°C and then washed them with PBS. Cells were reincubated in fresh medium at 37°C for 1 h to allow TMR-dextran to accumulate in lysosomes. The cells were then loaded with LysoSensor Green DND-189 for 15 min at 37°C, washed twice, and analyzed as described previously. When indicated, cells were stained with LysoSensor Green DND-153 (1 μM) according to a protocol similar to that used for LysoSensor Green DND-189. For flow cytometry, the macrophages were labeled as above, carefully removed from the culture plates, and immediately analyzed with a FACSCalibur flow cytometer (BD Biosciences).

Two laboratory strains of P. aeruginosa, ATCC 27853 and PAO1, were plated overnight on tryptic soy agar plates at 37°C, resuspended in tryptic soy broth at an OD of 0.25 at 550 nm, and incubated at 37°C for 1 h with shaking at 120 rpm. Bacteria were harvested in the early logarithmic growth phase and washed twice in RPMI 1640 supplemented with 1 mM HEPES (pH 7.4). Macrophages were infected with P. aeruginosa at a multiplicity of infection (MOI) of 100 (one macrophage was infected with 100 bacteria). For P. aeruginosa ATCC 27853 infection, synchronous infection conditions and an enhanced interaction between bacteria and host cells were achieved by a 2-min centrifugation (300 × g) of the bacteria onto the cells. The end of the centrifugation step was defined as the starting point of all infections. For P. aeruginosa PAO1 infection, bacteria were added without centrifugation. Bacterial CFUs were counted by serial dilution and plating on tryptic soy broth agar plates.

To determine intracellular ROS production, we loaded cells with 10 μM 2′,7′ dichlorodihydrofluorescein diacetate acetyl ester (H2DCFDA; Molecular Probes) for 30 min at 37°C. After incubation, cells were extensively washed with PBS and further incubated at 37°C for 10 min to allow cleavage of H2DCFDA to H2DCF by cellular esterases. Cells were left untreated or were treated with superoxide dismutase (SOD; 100 U/ml) and catalase (10 U/ml) for 10 min and infected with P. aeruginosa ATCC 27853 or PAO1 at an MOI of 100 for 1 h at 37°C. Postinfection, the mean fluorescence intensity was determined by flow cytometry (FACS-Calibur, BD Biosciences). The relative ROS production induced by P. aeruginosa, as indicated by the change in the fluorescence of 2′,7′-dichlorodihydrofluorescein diacetate (DCF), was calculated by using the following formula: ΔDCF fluorescence = F[PA]F[SOD + catalase + PA] (arbitrary unit) where F represents fluorescence intensity and PA represents P. aeruginosa.

Cells were infected with P. aeruginosa as above or left uninfected and were fixed in 2% paraformaldehyde/PBS for 10 min. To permeabilize cell membranes, we incubated cells with 0.1% Triton X-100/PBS for 10 min. We then washed the cells with PBS and incubated them for 30 min in PBS supplemented with 5% FCS to block nonspecific binding sites. Cells were washed again and incubated for 45 min with one of two primary Abs as indicated. The concentrations of these primary Abs were 1 μg/ml rat anti–lysosomal-associated membrane protein 1 (LAMP1) (BD Biosciences), 1 μg/ml mouse monoclonal anti-Cftr IgG (Millipore, Schwalbach, Germany), 1 μg/ml mouse monoclonal anti-ceramide IgM (Glycobiotech, Borstel, Germany), and 0.5 μg/ml monoclonal anti-gp91phox IgG (BD Biosciences). Cells were then washed in PBS with 0.05% Tween-20 and incubated for an additional 45 min with 0.5 μg/ml corresponding secondary Abs, namely, FITC–anti-rat IgG F(ab′)2 fragments, Cy3–anti-mouse IgG F(ab′)2 fragments, Cy3–anti-mouse IgM F(ab′)2 fragments, or FITC-labeled goat anti-mouse F(ab′)2 fragments, all from Jackson ImmunoResearch Laboratories (West Grove, PA) and diluted in PBS/5% FCS. After a final wash with PBS, cells were mounted on glass coverslips with Moviol. Control experiments were performed with irrelevant mouse Abs and secondary Abs. Control Abs did not substantially bind to the cells. Cells were examined on a Leica TCS SP confocal microscope (Leica Microsystems) equipped with a 100× oil objective, and images were analyzed with Leica Confocal Software (Leica Microsystems).

To determine bactericidal capability, we cultured macrophages in 96-well plates (105 cells per well) and infected them with P. aeruginosa strain ATCC 27853 or PAO1 opsonized in 5% mouse serum (MOI = 1). Phagocytosis of ATCC 27853 was synchronized by centrifugation at 300 × g for 5 min, and macrophages were infected for 20 min at 37°C. Infection with PAO1 bacteria was performed without centrifugation. Macrophages were then gently washed three times with sterile PBS for removal of extracellular or attached bacteria. Macrophages were further infected for 0 or 60 min at 37°C in the absence or presence of the NADPH oxidase inhibitor apocynin (100 μM). Postinfection, macrophages were incubated with 20 μl saponin (5 mg/ml) for 5 min for release of intracellular bacteria. Then 100 μl tryptose phosphate broth and 15 μl MTT in PBS (5 mg/ml) were added to each well, and the cells were incubated at 37°C for 4 h. Reactions were stopped by adding 100 μl isopropanol with 0.04 M HCl, and absorbance (A570nm) was measured at 570 nm with a microplate reader (BMG Labtech, Offenburg, Germany). To obtain the A570nm-CFU relationship, we incubated 103–105P. aeruginosa ATCC 27853 or PAO1 bacteria with 100 μl tryptose phosphate broth and 20 μl MTT in PBS (5 mg/ml) without macrophages for 4 h.

Macrophages were cultured in 24-well plates (5 × 105 cells per well in 1 ml RPMI1640 with 1 mM HEPES) and infected with the P. aeruginosa strains ATCC 27853 or PAO1 (MOI 100) for 2 h. KC and MIP-2 levels in the supernatants were determined by ELISA (R&D Systems, Wiesbaden-Nordenstadt, Germany) according to the manufacturer’s instructions.

Data are expressed as arithmetic means ± SD. Data obtained from multiple groups were tested using one-way ANOVA followed by Bonferroni post hoc t test. Significance between two groups (e.g., infected WT cells versus infected Cftr-deficient cells) was determined by Student t test as indicated in the figure legends. Values of p < 0.05 was considered statistically significant.

To determine whether Cftr deficiency affects lysosomal pH, we stained freshly isolated alveolar macrophages with LysoSensor Green DND-189, which accumulates in acidic organelles and exhibits green fluorescence. The results demonstrated that Cftr-deficient macrophages obtained from CftrMHH mice exhibit a much lower fluorescence intensity than do WT cells. This finding indicates that Cftr deficiency impairs the acidification of these vesicles (Fig. 1A). Bafilomycin is known to specifically inhibit vacuolar type H+-ATPase and, therefore, to elevate lysosomal pH (22). Pretreatment of cells with bafilomycin decreased LysoSensor fluorescence intensity in both WT and CftrMHH cells (Fig. 1A). To obtain a standard curve for LysoSensor Green DND-189 fluorescence, we permeabilized the cells with nigericin and incubated the cells that were previously stained with LysoSensor Green DND-189 in a calibration buffer with pH values ranging from 4.5–6.5. We found that the relative fluorescence of the LysoSensor Green DND-189 staining was linear over the investigated range of pH (Fig. 1B). A linear, pH-dependent decrease of intracellular LysoSensor Green DND-189 fluorescence was observed in both WT and CftrMHH cells. FACS analysis confirmed that bafilomycin decreased LysoSensor Green DND-189 fluorescence in WT macrophages (Fig. 1C).

FIGURE 1.

Cftr deficiency increases the pH of lysosomes. A, WT or Cftr-deficient (CFMHH) macrophages were treated with bafilomycin (100 nM) for 30 min or left untreated. They were then stained with LysoSensor Green DND-189 and analyzed by fluorescence microscopy. Representative fluorescence images or the mean ± SD from four independent experiments are shown. B, Macrophages were loaded with LysoSensor Green DND-189 and incubated with or without a calibration buffer adjusted to pH values ranging from 4.5–6.5. Relative LysoSensor Green DND-189 fluorescence of samples with calibration buffer was calculated as the percentage of samples with a pH of 4.5. The results were repeated three times with similar results. C, The panel displays a representative histogram produced by flow cytometric analysis of LysoSensor Green DND-189 fluorescence in WT macrophages with or without pretreatment with bafilomycin. The data are representative of those from three independent experiments. D and E, To determine which vesicles were stained with LysoSensor Green DND-189 and which with TMR-dextran, we loaded WT and Cftr-deficient macrophages with various concentrations of TMR-dextran and then stained them with LysoSensor Green DND-189. The results show that LysoSensor Green DND-189 also stains a vesicle population, which is not stained with TMR-dextran. Representative confocal fluorescence images from three independent experiments are shown.

FIGURE 1.

Cftr deficiency increases the pH of lysosomes. A, WT or Cftr-deficient (CFMHH) macrophages were treated with bafilomycin (100 nM) for 30 min or left untreated. They were then stained with LysoSensor Green DND-189 and analyzed by fluorescence microscopy. Representative fluorescence images or the mean ± SD from four independent experiments are shown. B, Macrophages were loaded with LysoSensor Green DND-189 and incubated with or without a calibration buffer adjusted to pH values ranging from 4.5–6.5. Relative LysoSensor Green DND-189 fluorescence of samples with calibration buffer was calculated as the percentage of samples with a pH of 4.5. The results were repeated three times with similar results. C, The panel displays a representative histogram produced by flow cytometric analysis of LysoSensor Green DND-189 fluorescence in WT macrophages with or without pretreatment with bafilomycin. The data are representative of those from three independent experiments. D and E, To determine which vesicles were stained with LysoSensor Green DND-189 and which with TMR-dextran, we loaded WT and Cftr-deficient macrophages with various concentrations of TMR-dextran and then stained them with LysoSensor Green DND-189. The results show that LysoSensor Green DND-189 also stains a vesicle population, which is not stained with TMR-dextran. Representative confocal fluorescence images from three independent experiments are shown.

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Next, we compared LysoSensor Green DND-189 staining of macrophages with TMR-dextran labeling of WT and CftrMHH macrophages. Cells were incubated with various concentrations of TMR-dextran and then stained with LysoSensor Green DND-189. The use of TMR-dextran at higher concentrations resulted in the staining of vesicles that did not stain with LysoSensor Green DND-189; at lower concentrations of TMR-dextran, all vesicles that stained with TMR-dextran also stained with LysoSensor Green DND 189, but not all LysoSensor Green DN189-positive vesicles were also positive for TMR-dextran (Fig. 1D, 1E). The higher concentrations of the weak base TMR-dextran very likely alkalinized the vesicles and thus prevented staining with LysoSensor Green DND-189 (Fig. 1D, 1E). These results indicate very clearly that LysoSensor Green DND-189 stains two populations of intracellular vesicles, one that is also stained by TMR-dextran and one that is not stained with TMR-dextran and probably consists of secretory lysosomes that are not stained with dyes that require internalization, such as TMR-dextran. Cftr deficiency substantially decreased the fluorescence intensity of LysoSensor Green DND-189 in these vesicles.

To exclude the possibility that Cftr-deficient cells have defects in the uptake or retention of LysoSensor probes, we stained 5 × 104 cells with LysoSensor Green DND-189, disrupted the cells in 200 μl pH calibration buffer (pH = 4 or 6.5) containing 1% Triton X-100, and then measured the total fluorescence intensity of the lysate with a fluorescence microplate reader (BMG Systems, Offenburg, Germany). No difference was found in the total fluorescence intensity of LysoSensor Green DND-189 between WT and Cftr-deficient cells either at an acidic pH of 4 or at a higher pH of 6.5 (Fig. 2A).

FIGURE 2.

Effect of Cftr deficiency on uptake and retention of LysoSensor probes and lysosome biogenesis. A, WT and Cftr-deficient (CFMHH) macrophages were loaded with LysoSensor Green DND-189 and lysed in pH calibration buffer (pH = 4.0 or 6.5) containing 1% Triton X-100. Total LysoSensor Green DND-189 fluorescence was then determined with a fluorescence microplate reader. Results are shown as the mean ± SD of 4 independent experiments. B, Macrophages were stained with TMR-dextran and LysoSensor Green DND-153, which is fluorescent at neutral pH. Representative confocal fluorescence images from three independent experiments are shown. C, Macrophages were fixed, permeabilized, and stained with FITC-labeled anti-LAMP1, Cy3-labeled anti-Cftr, or both. Representative fluorescence images from three independent experiments are shown.

FIGURE 2.

Effect of Cftr deficiency on uptake and retention of LysoSensor probes and lysosome biogenesis. A, WT and Cftr-deficient (CFMHH) macrophages were loaded with LysoSensor Green DND-189 and lysed in pH calibration buffer (pH = 4.0 or 6.5) containing 1% Triton X-100. Total LysoSensor Green DND-189 fluorescence was then determined with a fluorescence microplate reader. Results are shown as the mean ± SD of 4 independent experiments. B, Macrophages were stained with TMR-dextran and LysoSensor Green DND-153, which is fluorescent at neutral pH. Representative confocal fluorescence images from three independent experiments are shown. C, Macrophages were fixed, permeabilized, and stained with FITC-labeled anti-LAMP1, Cy3-labeled anti-Cftr, or both. Representative fluorescence images from three independent experiments are shown.

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We also stained the cells with LysoSensor Green DND-153, which exhibits bright fluorescence only at a neutral pH. The results demonstrated that at least some vesicles of Cftr-deficient macrophages exhibit a higher fluorescence intensity of LysoSensor Green DND-153 than do the vesicles of WT cells (Fig. 2B). These data indicate that Cftr deficiency results in pH changes in some vesicles rather than in an alteration in the uptake or retention of LysoSensor probes.

To determine whether Cftr resides in lysosomal membranes, we double-stained cells for Cftr and LAMP1. The results demonstrated that Cftr colocalizes with LAMP1 in some but not all vesicles of freshly isolated lung macrophages; this finding suggests that lysosomal membrane-associated Cftr directly contributes to the control of pH in lysosomes (Fig. 2C). No difference in LAMP1 staining was found between WT and Cftr-deficient macrophages, a finding suggesting that lysosome biogenesis is not changed by Cftr deficiency (Fig. 2C). The fact that Cftr-deficient cells do not stain with the anti-Cftr Abs indicates the specificity of the staining.

We have previously shown that Cftr-deficient epithelial cells accumulate ceramide (5). We suggested that the impaired acidification of vesicles, cellular compartments, or both may cause an imbalance in the activities of acid sphingomyelinase and acid ceramidase, and that this imbalance may result in the accumulation of ceramide (5). However, it is unknown whether ceramide also accumulates in Cftr-deficient macrophages.

To determine ceramide levels in macrophages, we permeabilized WT and Cftr-deficient alveolar macrophages obtained from CftrMHH mice and stained them with anti-ceramide Abs. We detected a marked accumulation of cellular ceramide in uninfected Cftr-deficient cells compared with WT cells (Fig. 3A). To confirm ceramide accumulation in Cftr-deficient macrophages, we performed DAG kinase assays that determine total cellular ceramide concentrations. We detected a substantial increase in ceramide concentrations in CftrMHH and CftrKO macrophages (Fig. 3B). Infection with P. aeruginosa increased intracellular ceramide staining in WT cells but had only a minimal effect on ceramide concentrations in CftrMHH macrophages (Fig. 3A).

FIGURE 3.

Cftr deficiency results in ceramide accumulation, clustering, and colocalization with gp91phox. A, Macrophages were infected with P. aeruginosa strain ATCC 27853. For detection of intracellular ceramide and gp91phox, cells were washed, fixed, permeabilized, and stained with Cy3-labeled ceramide and FITC-labeled gp91phox Abs. Shown are representative confocal fluorescence microscopic images of WT and Cftr-deficient (CftrMHH) cells that were either left uninfected or infected with P. aeruginosa (MOI = 100) for 15 min (n = 3). B, DAG kinase assays of total cellular ceramide contents demonstrate increased total levels of ceramide in Cftr-deficient macrophages compared with WT macrophages. Data are presented as the mean ± SD of results from three independent experiments. Significant differences of samples from Cftr-deficient compared with WT samples were determined by one-way ANOVA, followed by a Bonferroni post hoc t test and are indicated by an asterisk (* p < 0.05). C, Cftr deficiency impairs P. aeruginosa-induced clustering of gp91phox in ceramide-enriched membrane platforms. For detection of ceramide-enriched membrane platforms, macrophages were infected, fixed, and stained as in A, except that no permeabilization was performed because ceramide clusters would not be observed in permeabilized cells. Representative fluorescence images from three independent experiments are shown.

FIGURE 3.

Cftr deficiency results in ceramide accumulation, clustering, and colocalization with gp91phox. A, Macrophages were infected with P. aeruginosa strain ATCC 27853. For detection of intracellular ceramide and gp91phox, cells were washed, fixed, permeabilized, and stained with Cy3-labeled ceramide and FITC-labeled gp91phox Abs. Shown are representative confocal fluorescence microscopic images of WT and Cftr-deficient (CftrMHH) cells that were either left uninfected or infected with P. aeruginosa (MOI = 100) for 15 min (n = 3). B, DAG kinase assays of total cellular ceramide contents demonstrate increased total levels of ceramide in Cftr-deficient macrophages compared with WT macrophages. Data are presented as the mean ± SD of results from three independent experiments. Significant differences of samples from Cftr-deficient compared with WT samples were determined by one-way ANOVA, followed by a Bonferroni post hoc t test and are indicated by an asterisk (* p < 0.05). C, Cftr deficiency impairs P. aeruginosa-induced clustering of gp91phox in ceramide-enriched membrane platforms. For detection of ceramide-enriched membrane platforms, macrophages were infected, fixed, and stained as in A, except that no permeabilization was performed because ceramide clusters would not be observed in permeabilized cells. Representative fluorescence images from three independent experiments are shown.

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Previous studies from our group have demonstrated that the activation of acid sphingomyelinase and the release of ceramide on infection with P. aeruginosa trigger the release of ROS in macrophages (14). We therefore investigated whether P. aeruginosa infection induces colocalization of ceramide with gp91phox, an important membrane-bound subunit of NADPH oxidase, in vesicles of WT cells, and whether Cftr is involved in the colocalization of gp91phox and ceramide. The results show that P. aeruginosa triggers colocalization of gp91phox and ceramide in WT macrophages. Some degree of colocalization of gp91phox and ceramide was observed in CftrMHH macrophages before infection with P. aeruginosa (Fig. 3A). However, infection failed to induce further colocalization of gp91phox and ceramide.

Previous studies have shown that ceramide molecules form ceramide-enriched membrane platforms in the plasma membrane and that these ceramide molecules are crucially involved in signal transduction events that are controlled by ceramide (23, 24). To test whether P. aeruginosa infections trigger the formation of ceramide-enriched membrane domains in macrophages, we stained intact cells, namely, cells that were not permeabilized, with anti-ceramide Abs (Fig. 3C). The results show that P. aeruginosa induces massive clustering of ceramide in the plasma membrane of WT cells but that only very small ceramide-enriched membrane domains are observed in CftrMHH macrophages infected with P. aeruginosa (Fig. 3C). Costaining with anti-gp91phox Abs revealed that gp91phox is localized within ceramide-enriched membrane domains in WT macrophages (Fig. 3C). In contrast, we observed no increase in the colocalization of gp91phox and ceramide in CftrMHH macrophages on infection with P. aeruginosa (Fig. 3C; please note that cells in C are not permeabilized). However, Cftr deficiency alone resulted in some colocalization of ceramide and gp91phox in the plasma membrane already in uninfected cells.

The impaired colocalization of gp91phox and ceramide in Cftr-deficient macrophages suggests that Cftr is also involved in the regulation of the release of ROS on infection with P. aeruginosa. To examine ROS production by macrophages on infection with P. aeruginosa, we loaded cells with H2DCF (an ROS probe) and infected them with P. aeruginosa in the absence or presence of the ROS scavengers SOD and catalase. When the cells were not infected, there was no significant difference between WT and Cftr-deficient cells in the increase of DCF fluorescence, although ROS production was slightly increased in CftrMHH cells. Infection with P. aeruginosa ATCC 27853 or PAO1 induced a substantial increase in DCF fluorescence in WT macrophages including ROS production but not in CftrMHH macrophages (Fig. 4A).

FIGURE 4.

Cftr deficiency impairs the production of ROS and the bactericidal capability of macrophages. A, Macrophages were loaded with DCF, an ROS probe. The cells were then infected with P. aeruginosa ATCC 27853 or PAO1 strains (MOI = 100) for 1 h in the presence or absence of 100 U/ml SOD, an ROS scavenger, and 10 U/ml catalase. The change in DCF fluorescence was determined by using flow cytometry to measure the mean fluorescence intensity. Relative ROS production induced by P. aeruginosa, as indicated by the change in the fluorescence of DCF, was calculated by using the following formula: ΔDCF fluorescence = F[PA]F[SOD + catalase + PA] (arbitrary unit), where PA stands for P. aeruginosa, and F for fluorescence. Data are presented as the mean ± SD of results from four independent experiments. Significant differences between infected samples and uninfected controls were determined by one-way ANOVA followed by Bonferroni post hoc t test (*p < 0.05). Significant differences between infected WT cells and infected Cftr-deficient cells were determined by Student t test (Δ, p < 0.05). B, The survival of P. aeruginosa ATCC 27853 or PAO1 strains in macrophages postinfection for 60 min is displayed. As indicated, the macrophages were pretreated with apocynin (100 μM). Data are presented as the mean ± SD of results from three independent experiments. Significant differences between P. aeruginosa ATCC 27853-infected Cftr-deficient cells or WT cells treated with apocynin and WT controls were determined by one-way ANOVA followed by Bonferroni post hoc t test. Significant differences between P. aeruginosa PAO1-infected Cftr-deficient cells and WT controls were analyzed by Student t test. Significant differences (p < 0.05) are indicated by asterisks (*).

FIGURE 4.

Cftr deficiency impairs the production of ROS and the bactericidal capability of macrophages. A, Macrophages were loaded with DCF, an ROS probe. The cells were then infected with P. aeruginosa ATCC 27853 or PAO1 strains (MOI = 100) for 1 h in the presence or absence of 100 U/ml SOD, an ROS scavenger, and 10 U/ml catalase. The change in DCF fluorescence was determined by using flow cytometry to measure the mean fluorescence intensity. Relative ROS production induced by P. aeruginosa, as indicated by the change in the fluorescence of DCF, was calculated by using the following formula: ΔDCF fluorescence = F[PA]F[SOD + catalase + PA] (arbitrary unit), where PA stands for P. aeruginosa, and F for fluorescence. Data are presented as the mean ± SD of results from four independent experiments. Significant differences between infected samples and uninfected controls were determined by one-way ANOVA followed by Bonferroni post hoc t test (*p < 0.05). Significant differences between infected WT cells and infected Cftr-deficient cells were determined by Student t test (Δ, p < 0.05). B, The survival of P. aeruginosa ATCC 27853 or PAO1 strains in macrophages postinfection for 60 min is displayed. As indicated, the macrophages were pretreated with apocynin (100 μM). Data are presented as the mean ± SD of results from three independent experiments. Significant differences between P. aeruginosa ATCC 27853-infected Cftr-deficient cells or WT cells treated with apocynin and WT controls were determined by one-way ANOVA followed by Bonferroni post hoc t test. Significant differences between P. aeruginosa PAO1-infected Cftr-deficient cells and WT controls were analyzed by Student t test. Significant differences (p < 0.05) are indicated by asterisks (*).

Close modal

To determine whether the impaired release of ROS by Cftr-deficient macrophages postinfection with P. aeruginosa decreases the bactericidal capability of these macrophages, we incubated cells with P. aeruginosa at a low MOI (MOI = 1). WT macrophages eliminated ∼50% of the bacteria within 60 min when infected with P. aeruginosa ATCC 27853, whereas CftrMHH macrophages failed to kill the bacteria (Fig. 4B). Similar results were obtained with lung macrophages from CftrKO mice (Fig. 4B). The failure of Cftr-deficient macrophages to kill P. aeruginosa ATCC 27853 was mimicked by treating WT macrophages with the NADPH oxidase inhibitor apocynin (Fig. 4B). Similar results were found when WT or Cftr-deficient cells were infected with P. aeruginosa PAO1 (Fig. 4). Taken together, these findings indicate the importance of ROS for the killing of P. aeruginosa by macrophages.

Lung alveolar macrophages also play a crucial role in the production of cytokines during innate immune responses. Thus, we examined whether Cftr-deficient macrophages exhibit defects in cytokine production upon infection with P. aeruginosa. However, WT and Cftr-deficient macrophages produced similar amounts of KC and MIP2 during a 2-h infection with P. aeruginosa ATCC 27853 or PAO1 (Fig. 5A, 5B). These data suggest that the production of at least some cytokines (i.e., KC and MIP2) is not a consequence of Cftr deficiency in macrophages.

FIGURE 5.

Effects of Cftr deficiency on KC and MIP2 release by alveolar macrophages on infection with P. aeruginosa. Macrophages were infected with P. aeruginosa ATCC 27853 or PAO1 strains for 2 h. Cytokine levels of KC (A) and MIP2 (B) in the supernatants were measured with ELISA kits (R&D Systems). Significant differences between infected WT or Cftr-deficient cells and uninfected WT or Cftr-deficient controls were determined by one-way ANOVA followed by Bonferroni post hoc t test and are indicated by asterisks (*p < 0.05).

FIGURE 5.

Effects of Cftr deficiency on KC and MIP2 release by alveolar macrophages on infection with P. aeruginosa. Macrophages were infected with P. aeruginosa ATCC 27853 or PAO1 strains for 2 h. Cytokine levels of KC (A) and MIP2 (B) in the supernatants were measured with ELISA kits (R&D Systems). Significant differences between infected WT or Cftr-deficient cells and uninfected WT or Cftr-deficient controls were determined by one-way ANOVA followed by Bonferroni post hoc t test and are indicated by asterisks (*p < 0.05).

Close modal

In the current study, we demonstrate that freshly isolated lung macrophages contain at least two distinct acidic vesicle populations. Some, but not all of these vesicles exhibit a higher pH in Cftr-deficient macrophages than the corresponding vesicles in WT cells, as indicated by staining with LysoSensor Green DND-189 and DND-153.

The role of Cftr in the regulation of vesicular pH is controversial. Barasch and Al-Awqati demonstrated the alkalinization of trans-Golgi and mannose-6-phosphate–positive vesicles and prelysosomes in Cftr-deficient cells (3). These findings were confirmed by the results of several studies that demonstrated the alkalinization of lysosomes and phagosomes in alveolar macrophages (3, 4, 7). However, other reports have failed to detect a role of Cftr in the regulation of pH in vesicles that form after internalization (10, 11). These studies determined the pH in TMR-dextran–positive endosomes or vesicles that contain internalized Ab-labeled or fluorescent Cftr molecules (10, 11). Dextran-containing endosomes are directly targeted to a subpopulation of lysosomes, and these endosomes are distinct from mannose-6-phosphate–positive vesicles and do not mix with secretory lysosomes (25). The discrepancies in findings about the role of Cftr in the regulation of vesicular pH could be caused by the fact that the reported studies used different cell systems and different techniques. Moreover, different types of vesicles express different sets of ion channels and pumps (26), and this fact very likely explains the differences in the effects of Cftr deficiency on the pH of a particular population of vesicles. The findings of the current study confirm the notion that Cftr is involved in the regulation of the pH in a subset of intracellular vesicles, most likely secretory lysosomes.

Changes in the pH of secretory lysosomes may result in a relative overactivity of acid sphingomyelinase, which is still active at a pH of 6.0, in comparison with that of acid ceramidase, which is almost inactive at pH 6.0; this overactivity might be followed by a slow accumulation of ceramide in these vesicles of Cftr-deficient cells (5). Indeed, we observed a marked chronic accumulation of ceramide in Cftr-deficient alveolar macrophages. In addition, Cftr deficiency prevented the acute and rapid activation of acid sphingomyelinase observed in WT cells postinfection with P. aeruginosa (15), thereby preventing the acute release of ceramide on infection. On the basis of these observations, we assume that the LysoSensor Green DND-189–positive lysosomes are secretory lysosomes containing the signaling pool of acid sphingomyelinase, that their vesicular pH is regulated by Cftr and that the increase in the pH of secretory lysosomes prevents activation of the acid sphingomyelinase. In contrast, the lysosomal form of acid sphingomyelinase, which mediates the physiological steady-state metabolism of sphingomyelin, does not seem to be affected in Cftr-deficient cells, because classical lysosomes in Cftr-deficient cells do not seem to exhibit an accumulation of sphingomyelin.

We further explored whether the influence of Cftr deficiency is linked to alterations in lysosomal biogenesis and transport. We did not detect a difference in staining with LAMP1 between WT and Cftr-deficient macrophages; this finding suggests that lysosome biogenesis is not altered in the absence of Cftr. The finding that Cftr colocalizes with LAMP1 further supports the hypothesis that Cftr resides in the lysosomal membrane and is directly involved in the acidification of lysosomes. Finally, Cftr deficiency exerted no effect on TMR-dextran staining. Because TMR-dextran is internalized by endocytosis and released by vesicular exocytosis, TMR-dextran staining in the cells reflects equilibrium between endocytosis and vesicular exocytosis. Thus, our findings indicate that the formation of endocytotic and secretory vesicles is not altered by Cftr deficiency. Taken together, our findings strongly suggest that Cftr is directly involved in the regulation of pH in a specific subpopulation of lysosomes but is not involved in lysosomal biogenesis or the formation of endocytic and secretory vesicles in lung alveolar macrophages.

However, it is also possible that Cftr deficiency affects the pH on the extracellular leaflet of the cell membrane and thereby alters the balance between acid sphingomyelinase and acid ceramidase. Moreover, it is possible that Cftr deficiency prevents the acute activation of acid sphingomyelinase by P. aeruginosa because it alters the lipid content of secretory lysosomes. Previous studies by Tabas and colleagues demonstrated that the lipid composition of membranes has a substantial impact on the activation of acid sphingomyelinase (27). It remains to be determined whether alterations of other membrane lipids in Cftr-deficient cells contribute to the defect of acid sphingomyelinase activation in these cells.

Previous studies have demonstrated that acid sphingomyelinase plays a role in the activation of NADPH oxidase by TNF-α, CD95, and infection with P. aeruginosa (14, 28, 29). Activation of the acid sphingomyelinase seems to occur in secretory lysosomes that fuse with the cell membrane (23, 29). Ceramide then forms ceramide-enriched membrane platforms in the plasma membrane; these platforms serve the clustering and activation of NADPH oxidase and, thus, the release of ROS (14). Studies using cells lacking acid sphingomyelinase showed that acid sphingomyelinase mediates the activation of NADPH oxidase and the release of ROS on infection with P. aeruginosa (14).

In the current study, we demonstrated that P. aeruginosa-induced ROS release is abolished in Cftr-deficient macrophages. This impairment in ROS release by Cftr-deficient macrophages also decreases the bactericidal capability of these cells against P. aeruginosa. In contrast, Cftr deficiency did not affect the cytokine release response by these lung macrophages, a finding that is consistent with those of previous studies showing that Cftr is not involved in the production of cytokines (KC and MIP2) by LPS-stimulated macrophages (30). This situation is in marked contrast to that in Cftr-deficient epithelial cells, which show a marked increase in the constitutive release of proinflammatory cytokines, such as IL-1 and IL-8 (5, 31, 32).

At present, the role of macrophages in CF is poorly defined. Although several studies have demonstrated that Cftr-deficient macrophages exhibit a defect in the intracellular killing of P. aeruginosa (4, 7), the role of macrophages in acute pulmonary infections with P. aeruginosa in vivo is still open (33). A recent study demonstrated that in particular neutrophils are essential in eliminating P. aeruginosa from infected airways (33). Thus, macrophages might be important in regulating pathways which enhance inflammation including amplification of neutrophil recruitment. It is also possible that the inability of Cftr-deficient macrophages to kill sufficient numbers of P. aeruginosa bacteria contributes to the initial sensitivity of CF patients (and mice) to infections with P. aeruginosa, a situation that does not apply to WT mice. This defect in Cftr-deficient macrophages may result in a slow removal of the bacteria from alveoli, a longer persistence of the bacteria in the lung, and finally the growth of the bacteria in the lung. It could be speculated that the survival of P. aeruginosa in Cftr-deficient macrophages may protect the pathogens from the neutrophil attack in CF. However, determining whether this hypothesis is true will require future studies that are beyond the focus of the current study; such studies will probably use floxed Cftr-deficient mice with a specific deficiency of Cftr in macrophages to exactly define the function of alveolar macrophages in CF.

In summary, we have demonstrated that Cftr deficiency results in a chronic accumulation of ceramide in macrophages but prevents the acute release of ceramide on infection of these cells with P. aeruginosa. This defect results in a failure to stimulate the activation of NADPH oxidase and, thus, an inability to release ROS and to kill P. aeruginosa. These studies link Cftr deficiency, via acid sphingomyelinase and ceramide, to the control of the release of ROS and the killing of P. aeruginosa.

We thank Siegfried Moyer for administrative and graphical assistance.

Disclosures The authors have no financial conflicts of interest.

This work was supported by Deutsche Forschungsgemeinschaft Grant Gu 335-16/1.

Abbreviations used in this paper:

ATPase

adenosine triphosphatase

CF

cystic fibrosis

CFTR

cystic fibrosis transmembrane conductance regulator

DAG

1,2-diacylglycerol

DCF

2′,7′-dichlorodihydrofluorescein diacetate

DTPA

diethylenetriaminepentaacetic acid

H2DCFDA

2′,7′-dichlorodihydrofluorescein diacetate acetyl ester

LAMP1

lysosome-associated membrane protein 1

MOI

multiplicity of infection

ROS

reactive oxygen species

SOD

superoxide dismutase

TMR

tetramethylrhodamine

WT

wild-type.

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