Dysfunctional cystic fibrosis transmembrane conductance regulator inhibits phagocytosis of apoptotic cells with proinflammatory consequences

Abstract

Cystic fibrosis (CF) is caused by mutated CF transmembrane conductance regulator (CFTR) and is characterized by robust airway inflammation and accumulation of apoptotic cells. Phagocytosis of apoptotic cells (efferocytosis) is a pivotal regulator of inflammation, because it prevents postapoptotic necrosis and actively suppresses release of a variety of proinflammatory mediators, including IL-8. Because CF is associated with accumulation of apoptotic cells, inappropriate levels of IL-8, and robust inflammation, we sought to determine whether CFTR deficiency specifically impairs efferocytosis and its regulation of inflammatory mediator release. Here we show that CFTR deficiency directly interferes with efferocytosis by airway epithelium, an effect that is not due to altered binding of apoptotic cells to epithelial cells or altered expression of efferocytosis receptors. In contrast, expression of RhoA, a known negative regulator of efferocytosis, is substantially increased in CFTR-deficient cells, and inhibitors of RhoA or its downstream effector Rho kinase normalize efferocytosis in these cells. Impaired efferocytosis appears to be mediated through an amiloride-sensitive ion channel, because amiloride restores phagocytic competency in CFTR-deficient cells. Finally, ineffective efferocytosis in CFTR-deficient cells appears to have proinflammatory consequences, because apoptotic cells enhance IL-8 release by these cells, but not by wild-type controls. Therefore, in CF, dysregulated efferocytosis may lead to accumulation of apoptotic cells and impaired regulation of the inflammatory response and, ultimately, may suggest a new therapeutic target.

early, intense, and unrelenting inflammation dominates the pathobiology of cystic fibrosis (CF) lung disease in the presence and absence of detectable infection (26); this central observation suggests that CF transmembrane conductance regulator (CFTR), a Cl⁻ channel that is the genetic cause of CF, is a pivotal regulator of the innate immune response and, ultimately, lung homeostasis.

Pathogenic microorganisms play a clear and critical role in the development of CF lung disease, especially over a lifetime. Yet, in vitro, ex vivo, and in vivo studies increasingly suggest that CFTR deficiency also causes a primary dysregulation of the inflammatory response. For example, dysfunctional CFTR increases NFκB activity and enhances inflammatory mediator release (e.g., IL-8) by epithelial cells (12), possibly due to overload of the endoplasmic reticulum (12), enhanced oxidative stress, altered processing of IκBβ (56), impaired signaling by anti-inflammatory mediators (24), an exaggerated response to hypertonicity, or impaired lipoxin metabolism (22). Naive human tracheal grafts from CF fetuses spontaneously develop inflammation in the absence of infection and respond excessively to a bacterial challenge (51). When CFTR-deficient mice are challenged intratracheally with Pseudomonas aeruginosa, they mount an enhanced inflammatory response (17). Apoptotic inflammatory cells accumulate in the airways of young adults with CF, in part through ineffective removal, suggesting that failed phagocytosis may contribute to ongoing airway inflammation (53).

Efferocytosis, or phagocytosis of apoptotic cells, is a form of stimulated macropinocytosis that is integrally involved with regulation of the inflammatory response and lung homeostasis (18). Apoptotic cells flag themselves for recognition and uptake into phagocytes by expressing surface ligands, in particular phosphatidylserine (PS) (18, 54). A variety of structures are involved in the removal process (18, 20, 36, 41, 42, 54), especially those that recognize PS and members of the ATP-binding cassette (ABC) protein family, including ced-7 (61), ABC-A1 (32), and ABC-A7 (21). The involvement of ced-7, ABC-A1, and ABC-A7 is particularly noteworthy, because CFTR is also a member of this family of transporters, and efferocytosis is impaired in CF (53). Efferocytosis is tightly regulated by the Rho family of GTPases, in that it is stimulated by Rac-1 and Cdc42 (19, 31) and inhibited by RhoA and its downstream effector Rho kinase (52).

The net effect of efferocytosis is anti-inflammatory, because dying cells are removed before they undergo postapoptotic necrosis and because recognition of PS induces release of anti-inflammatory mediators, such as transforming growth factor-β1 (TGFβ1) (18, 54). These anti-inflammatory signals actively suppress IL-8 and other proinflammatory mediators (18, 54) that appear to be central in CF disease pathogenesis (26). In addition, efferocytosis causes release of growth factors and antiproteases that would be expected to contribute to tissue repair and counteract protease-antiprotease imbalance (18, 54).

Because CF is associated with accumulation of apoptotic cells, ineffective efferocytosis, and robust inflammation, we sought to determine whether CFTR deficiency specifically impairs efferocytosis and its regulation of inflammatory mediator release. Results indicate that CFTR deficiency directly interferes with efferocytosis by airway epithelial cells and that the consequences are proinflammatory. CFTR deficiency appears to mediate these processes, in part, through a RhoA-dependent and amiloride-sensitive mechanism.

MATERIALS AND METHODS

Materials.

Mouse monoclonal anti-human CD36 IgM, anti-human CD44 IgG1, anti-human β₃-integrin IgG1, biotin- and non-biotin-conjugated anti-human CD59 IgG2a, anti-human CD32 [Fcγ receptor IIa (FcγRIIa)] IgG2b, and mouse monoclonal IgG1, IgG2a, IgG2b, IgM, and κ-isotype control antibodies were obtained from BD Biosciences PharMingen (San Diego, CA); mouse monoclonal anti-human α_vβ₅-integrin IgG1 and mouse monoclonal anti-human α_V-integrin IgG1 antibodies from Chemicon International (Temecula, CA); chicken polyclonal anti-human calreticulin IgY antibody from Affinity BioReagents (Golden, CO); and mouse monoclonal anti-human α-chain CD91 IgG1 antibody from American Diagnostica (Greenwich, CT). Mouse monoclonal anti-human PS recognition structure IgM was prepared in this laboratory as previously described (53). Cy3 goat IgG anti-mouse IgM and Cy3 goat IgG anti-mouse IgG secondary antibodies were purchased from Jackson ImmunoResearch Laboratories (West Grove, PA); human TNFα from R & D Systems (Minneapolis, MN); and human IL-1β from Sigma-Aldrich (St. Louis, MO).

Human subjects.

The study was approved by and performed in accordance with the ethical standards of the Institutional Review Board on Human Experimentation at the University or Toronto and University of Colorado Health Sciences Center. Written informed consent was obtained from each subject.

Experimental animals.

Experiments were carried out according to protocols approved by the Animal Care and Use Committee at the Hospital for Sick Children (Toronto, ON, Canada). Long-surviving, liquid-fed, out-bred Cftr^m1UNC/Cftr^m1UNC knockout (CFTR^−/−) mice (25) and their littermates Cftr^+/+ wild-type (CFTR^+/+ control) mice (6–8 wk old) were utilized in the study. CFTR^+/+ mice were also fed a liquid diet during the experimental protocol to minimize differences that could be attributed to diet or nutritional status. Genotyping was done as previously described (25), and only homozygotes were used. CFTR^+/+ or CFTR^−/− mice were housed in a pathogen-free area in sterile microisolator cages until the time of experimentation. Littermate wild-type control mice were maintained under identical conditions.

Primary cells and cell lines.

Primary human airway epithelial cells were isolated as previously described (23) from the large airways of surgical explants from nine nondiseased donors and nine CF patients (8 CF patients were homozygous for the ΔF508 CFTR mutation, and 1 was compound heterozygous for ΔF508:1717-1G>A). Primary epithelial cells were cultured in bronchial epithelial cell growth medium (BEGM, Cambrex, East Rutherford, NJ). Human neutrophils were isolated from normal volunteers, as described elsewhere (53).

IB3-1 epithelial cells were kindly provided by Dr. Pam Zeitlin (Johns Hopkins School of Medicine, Baltimore, MD). IB3-1 cells are an adeno-12-SV40 immortalized bronchial epithelial cell line that was originally developed from a CF patient who was compound heterozygous for ΔF508:W1282X. C38 cells were developed from IB3-1 cells that were stably transfected with a truncated wild-type human CFTR. C38 and IB3-1 cells were grown in LHC-8 medium (Biosource International, Camarillo, CA) supplemented with antibiotics and FBS.

Jurkat T cells, a human T cell leukemia line, were obtained from American Type Culture Collection (Manassas, VA).

Induction of apoptosis.

Jurkat cells were exposed to ultraviolet irradiation at 312 nm for 10 min and cultured for 3.5 h. Jurkat cells were ∼70% apoptotic by nuclear condensation at the time of experimentation, as described previously (43).

In vitro phagocytosis and binding assays.

Phagocytosis assays were performed as described previously with modifications (53). Briefly, 1 × 10⁵ epithelial cells were seeded into 24-well plates (Fisher Scientific, Pittsburgh, PA) onto baked coverslips (Fisher Scientific), cultured in complete medium, and allowed to double over the next 24 h. In the case of primary epithelial cells, the coverslips were precoated with type IV human placental collagen (Sigma-Aldrich). Epithelial cells were then cocultured with 2.5 × 10⁶ apoptotic Jurkat cells suspended in 500 μl of X-Vivo medium (Cambrex Bio Science, Walkersville, MD) at 37°C in 5% CO₂ in the absence of human serum for 4 h. In some experiments, 8 × 10⁶ polystyrene beads (2–8 μm; Bangs Laboratories, Fishers, IN) were cocultured with epithelial cells at 37°C in 5% CO₂ in the absence of human serum for 4 h. The CFTR channel blocker CFTR_inh 172 (Sigma-Aldrich) or DMSO vehicle was added during the coculture period in some experiments. In other experiments, IB3-1 cells were cultured for 18 h before coculture at 37°C or 26°C (10, 12, 47). After coculture, epithelial cells were gently washed with PBS (Fisher Scientific) for removal of uningested cells or beads, fixed, and stained with modified Wright's Giemsa (Fisher Scientific). Phagocytosis and binding were determined by visual inspection of samples and expressed as phagocytic index (PI) or binding index (BI) (53). We considered apoptotic cells to be ingested if ≤20% of their margins extended beyond the edge of the epithelial cell and if they were within the optical plane of the epithelial cell. Most ingested apoptotic cells showed signs of digestion, such as blurring of the cell border or cell contents, and some were surrounded by the clear rim of a phagosome. We considered all apoptotic cells in contact with epithelial cells to be bound if they did not meet these criteria. These methods are subjective and could over- or underestimate apoptotic cell ingestions, but they correlated with the erythrocyte ingestion assay (see below). Each condition was tested in duplicate, and ≥400 epithelial cells per coverslip were counted blindly.

In vivo phagocytosis assays.

Phagocytosis assays were performed as described previously (38, 49, 55). Briefly, apoptotic human neutrophils (10 × 10⁶) suspended in 50 μl of PBS were instilled intratracheally through a modified animal feeding needle (Fisher Scientific). Whole lung lavage was performed with 5 ml of ice-cold PBS 30 min following intratracheal instillation of apoptotic cells. A cytospin (Thermo Electron, Pittsburgh, PA) slide of lung lavage was stained with modified Wright's Giemsa (Fisher Scientific) and evaluated for macrophage phagocytosis of apoptotic cells.

Erythrocyte ingestion assay.

Erythrocyte ingestion was assayed as previously described with modifications (19, 55). Normal human erythrocytes (E) were biotinylated with EZ-Link Sulfo-NHS-LC-Biotin (Pierce, Rockford, IL). Biotinylated E were then coated with streptavidin (Sigma-Aldrich) and linked to a biotinylated anti-human CD59 antibody (E-anti-CD59). Proper construction of E-anti-CD59 was confirmed by fluorescence-activated cell sorting (FACS) analysis using the appropriate fluorochrome-conjugated antibody. In some cases, E or E-anti-CD59 was stimulated with 250 nM ionomycin for 15 min to induce externalization of PS; these cells were designated E(PS) or E(PS)-anti-CD59. Externalization of PS was confirmed with annexin V staining. Modified and unmodified E were added to epithelial cells in duplicate wells at a 35:1 ratio and allowed to incubate at 37°C for 4 h. Unbound E were washed away with PBS. Uningested E were lysed by addition of deionized H₂O for 10 s in one of the two duplicate wells. Cells were fixed with 0.75% glutaraldehyde (Sigma-Aldrich), stained with dianisidine-H₂O₂ (Sigma-Aldrich), and counterstained with Wright's stain. Five hundred cells were scored using light microscopy to quantify binding (hypotonic lysis) and engulfment (after hypotonic lysis).

Fcγ receptor transfection.

C38 and IB3-1 epithelial cell lines were transfected with full-length human FcγRIIa in pCDNA3.1/GS (Invitrogen, Carlsbad, CA) using Lipofectamine 2000 reagent (Invitrogen) according to the manufacturer's instructions. Briefly, 5 × 10⁴ cells were seeded onto sterile coverslips in 24-well plates 24 h before transfection. Epithelial cells were then transfected with 0.4 μg of FcγRIIa/pCDNA 3.1/GS, 0.4 μg of pCDNA3.1/GS (empty vector), or Lipofectamine 2000 reagent (vehicle) in LHC-8 medium without serum at 37°C in 5% CO₂ for 3 h. After 48 h, FcγRIIa transfections were confirmed by FACS analysis using the appropriate primary and secondary antibodies, and transfection efficiency was ∼20% (data not shown). Transfected epithelial cells were cocultured for 4 h with 7.5 × 10⁶ human erythrocytes optimally coated (confirmed by FACS analysis) with anti-human erythrocyte rabbit IgG fraction (ICN Pharmaceuticals, Aurora, OH). Unbound E were washed away, hypotonically lysed in half of the wells, fixed with 0.75% glutaraldehyde, stained with dianisidine-H₂O₂, and counterstained with Wright's stain. Five hundred cells were scored using light microscopy to quantify binding (hypotonic lysis) and engulfment (after hypotonic lysis).

Inflammatory mediator experiments.

CFTR^+/+ and CFTR^−/− primary airway epithelial cells (3 × 10⁵) or C38 and IB3-1 epithelial cell lines (3 × 10⁵) were seeded onto six-well dishes and cultured in complete medium 24 h before the experiment; in the case of primary epithelial cells, the dishes were coated with collagen. Unstimulated, TNFα-stimulated (220 U/ml), and IL-1β-stimulated (50 ng/ml) epithelial cells were then cultured in the presence or absence of the indicated apoptotic cells in 1 ml of X-Vivo medium without FCS for 20 h. Some of these experiments were performed in the presence or absence of the Rho kinase inhibitor Y-27632 (Sigma-Aldrich). Supernatants were collected, assayed for IL-8 and TGFβ1, and corrected for cell numbers (ElisaTech, Aurora, CO).

RhoA activity assays and Western blots.

C38 and IB3-1 epithelial cell (5 × 10⁶) lysates were incubated in the presence or absence of GTPγS and then incubated with glutathione S-transferase (GST)-tagged Rho-binding domain of rhotekin bound to glutathione-Sepharose beads for 1 h at 4°C according to the manufacturer's instructions (Cytoskeleton, Denver, CO). Beads were washed, and binding of active RhoA (RhoA-GTP) in the lysates from the beads was analyzed by immunoblotting for RhoA. To confirm equal loading of proteins, 40 μl of total cell lysates for each sample were blotted for β-actin (Cell Signaling) after binding to rhotekin-bound glutathione-Sepharose beads. Western blotting for total RhoA was performed on cell lysates only.

Fluorescent staining for actin stress fibers.

C38 and IB3-1 cells were plated on glass coverslips in 24-well plates at 5 × 10⁵ cells per well 24 h before experimentation. Epithelial cells were then treated with complete medium, serum starved for 6 h with DMEM supplemented with 0.5% BSA for 6 h, or stimulated with 20 μM lysophosphatidic acid (LPA) for 10 min. Epithelial cells were then washed three times with PBS, fixed with 4% paraformaldehyde on ice for 20 min, permeabilized with 0.1% Triton X-100 in PBS at room temperature for 15 min, and stained with Alexa Fluor 488-phalloidin (Invitrogen) for 30 min at room temperature. Coverslips were mounted with 90% glycerol containing Hoechst 33258 (Sigma-Aldrich) at 0.5 μg/ml. Actin stress fibers were then imaged using a fluorescence microscope (Eclipse E800, Nikon).

Statistics.

Values are means ± SE. Means were analyzed using ANOVA for multiple comparisons; when ANOVA indicated significance, the Tukey-Kramer honestly significantly difference test for all pairs was used to compare groups. For all other experiments in which two conditions were compared, a Student's t-test assuming equal variance was used. All data were analyzed using JMP (version 3) statistical software for the Macintosh (SAS Institute, Cary, NC).

RESULTS

CFTR regulates epithelial cell efferocytosis in vitro.

Primary airway epithelial cells undergo spontaneous apoptosis in culture and are ingested by neighboring epithelial cells (Fig. 1, A and B). We found it to be particularly noteworthy that “nonprofessional” phagocytes (e.g., airway epithelial cells) were capable of ingesting dead cells of nearly equivalent size into large, spacious phagosomes, similar to ingestion of dead cells in efferocytosis by “professional” phagocytes, such as macrophages (Fig. 1, C and D). When spontaneous ingestions by primary CFTR^+/+ and CFTR^−/− epithelial cells were assessed, fewer ingestions were detected in CFTR^−/− epithelial cells (Fig. 1E), suggesting that CFTR regulates the rate of spontaneous apoptosis or efferocytosis.

Fig. 1.

CFTR deficiency is associated with fewer spontaneous efferocytosis events in vitro. Representative photomicrographs show spontaneous ingestion of apoptotic cells into spacious phagosomes by neighboring primary human airway epithelial cells (A and B) and human alveolar macrophages (C and D). Arrows indicate ingested apoptotic cells. E: quantitation of spontaneously ingested apoptotic cells by CFTR^+/+ and CFTR^−/− primary human airway epithelial cells [phagocytic index (PI)]. Values are means ± SE for 4 replicates per group. *Significantly different from CFTR^+/+ cells (P < 0.05).

To determine whether CFTR was involved with regulation of efferocytosis, primary CFTR^+/+ and CFTR^−/− epithelial cells were cocultured with apoptotic Jurkat T cells, and ingestion/binding was quantified (Fig. 2, A and B). Epithelial cells were also evaluated without coculture to determine the baseline level of spontaneous ingestion and binding. CFTR deficiency suppressed efferocytosis of Jurkat T cells in primary epithelial cells (Fig. 2C) but had no effect on binding (Fig. 2D). Similarly, efferocytosis, but not binding, of apoptotic Jurkat T cells was decreased in a CFTR^−/− airway epithelial cell line (IB3-1) compared with its CFTR-corrected control (C38) cell line (Fig. 2, E and F). These findings strongly suggest that CFTR plays a role in efferocytosis by airway epithelial cells.

Fig. 2.

CFTR deficiency decreases efferocytosis. A and B: representative photomicrographs showing ingestion of apoptotic Jurkat T cells by primary human airway epithelial cells. Arrows indicate ingested apoptotic cells. C and D: CFTR^+/+ and CFTR^−/− primary airway epithelial cells were cocultured with no cells or with apoptotic Jurkat T cells and analyzed for PI or binding index (BI). Values are means ± SE for 5–7 replicates per group. *Significantly different from identically treated CFTR^+/+ cells (P < 0.05). E and F: C38 and IB3-1 epithelial cells were cocultured with no cells or apoptotic Jurkat T cells for 4 h and analyzed for PI or BI. Values are means ± SE for 3 replicates per group. *Significantly different from identically treated C38 cells (P < 0.05). G: PI in CFTR^+/+ and CFTR^−/− mice instilled intratracheally with apoptotic polymorphonuclear leukocytes (neutrophils). After 30 min, whole lung lavage was performed, and alveolar macrophage PI was determined. Values are means ± SE for 5 animals per group.

CFTR is also expressed in alveolar macrophages and has been shown to affect functions such as lysosomal acidification (11). To address whether CFTR also regulates efferocytosis by alveolar macrophages in vivo, CFTR^+/+ and CFTR^−/− mice were challenged intratracheally with exogenous apoptotic neutrophils, and clearance was assessed in bronchoalveolar lavage. CFTR deficiency did not affect the alveolar macrophage PI (Fig. 2G), indicating that CFTR does not regulate efferocytosis by alveolar macrophages in the same way that it does for epithelial cells.

The effects of CFTR may be dependent or independent of its Cl⁻ channel function. To address this issue, apoptotic cell clearance experiments were performed with CFTR-sufficient C38 epithelial cells in the presence and absence of the CFTR channel blocker CFTR_inh 172 (Fig. 3A). Neither 5 nor 10 μM CFTR_inh 172 had a negative effect on efferocytosis, suggesting that CFTR's channel regulatory function had no role in efferocytosis. In contrast, a tendency toward a partial correction of the efferocytosis defect was observed in CFTR-deficient IB3-1 cells cultured at 26°C for 18 h (Fig. 3B).

Fig. 3.

CFTR ion channel function is not required for regulation of efferocytosis. A: to determine whether CFTR's ion channel function was required for regulation of efferocytosis by epithelial cells, C38 epithelial cells were cocultured with no cells or apoptotic Jurkat T cells for 4 h in the presence of DMSO or the specific CFTR channel inhibitor CFTR_inh 172 (5 and 10 μM). Values are means ± SE for 4 replicates per group. *Significantly different from untreated C38 cells cocultured with apoptotic cells (P < 0.05). B: culture of epithelial cells at low temperature increases translocation of the ΔF508 CFTR mutation to membrane, resulting in increased function in primary epithelial cells and IB3-1 cells (10, 12, 47). Efferocytosis was studied in IB3-1 cells cultured at 37°C or 26°C for 18 h and then cocultured with Jurkat T cells for 4 h at 37°C. Culture at low temperature tended to increase efferocytosis by IB3-1 cells. Values are means ± SE for 5 replicates per group. *Significantly different from 37°C (P = 0.08).

CFTR deficiency does not affect expression of efferocytosis recognition structures but does inhibit ingestion through a PS-mediated mechanism and FcγRIIa.

We examined the effect of CFTR on the expression of several key efferocytosis recognition structures by flow cytometry to address how CFTR might regulate efferocytosis. C38 and IB3-1 cells expressed equal amounts of calreticulin, CD44, CD36, and α_vβ₃- and α_vβ₅-integrins (data not shown). The low-density lipoprotein receptor-related protein (LRP or CD91) could not be detected on either cell line (data not shown). These data suggest that the effect of CFTR on efferocytosis was not due to differential receptor expression but do not rule out the possibility that receptor affinity may be impaired.

Receptor function was examined using an erythrocyte ingestion system that allows assessment of ligand-specific binding and uptake (Fig. 4, A and B) (19, 55). Because externalized PS is among the best-characterized “eat-me” signals on the apoptotic cell surface (18), the erythrocyte ingestion system was used to evaluate the effect of CFTR on PS-mediated phagocytosis. In this system, the presence of PS alone on the erythrocyte surface is not sufficient to drive ingestion. However, when PS-expressing erythrocytes are tethered to the phagocyte surface via binding to CD59, they are readily ingested (19).

Fig. 4.

CFTR deficiency inhibits phagocytosis mediated by the phosphatidylserine (PS) receptor and the Fcγ receptor IIa (FcγRIIa) but does not significantly affect latex bead ingestion. A and B: representative photomicrographs showing ingestion (uptake) and binding of E(PS)-αCD59 by C38 epithelial cells. C and D: PI (after hypotonic lysis) and BI (before hypotonic lysis) of various erythrocyte (E) targets cocultured with C38 and IB3-1 epithelial cells. Values are means ± SE for 5–6 replicates per group. *Significantly different from identically treated C38 cells (P < 0.05). E and F: PI and BI of IgG-coated erythrocytes cocultured with C38 or IB3-1 epithelial cells that had been transfected with vehicle, empty vector, or FcγRIIa. Values are means ± SE for 3 replicates per group. *Significantly different from identically treated C38 cells (P < 0.05).

We constructed erythrocyte targets (E) by linking them with a biotinylated anti-CD59 antibody (E-anti-CD59). We chose CD59 (membrane attack complex inhibitor), because, as the tethering target, it is equally present on C38 and IB3-1 cells (data not shown) and it is not known to be involved with any form of phagocytosis. E-anti-CD59 bound to epithelial cells without being ingested (Fig. 4, C and D). We also treated some E-anti-CD59 with the calcium ionophore ionomycin to cause membrane scrambling and PS externalization (30). [E(PS)-anti-CD59] were ingested through a PS-recognition mechanism (19). In these experiments, CFTR deficiency markedly inhibited PS-mediated ingestion (Fig. 4C) but had no effect on binding (Fig. 4D). These results mirror those for efferocytosis in Figs. 1 and 2.

We next sought to determine whether defective phagocytosis conferred by CFTR deficiency is specific to efferocytosis or could be generalized to other forms of phagocytosis, such as by the Fcγ receptor or latex beads. Epithelial cells do not express FcγRIIa, but FcγRIIa-transfected epithelium can ingest IgG-opsonized erythrocytes (13). Human FcγRIIa was transfected into C38 and IB3-1 cells, and uptake of IgG-opsonized erythrocytes was assessed. FcγRIIa transfection efficiency was ∼20% in C38 and IB3-1 cells (data not shown). Similar to efferocytosis and PS-mediated uptake, CFTR deficiency inhibited ingestion through FcγRIIa but had no effect on binding (Fig. 4, E and F). In contrast, CFTR had no significant effect on latex bead uptake (data not shown). Together, these data suggest that CFTR deficiency causes a broad defect in phagocytosis that is not limited to efferocytosis.

CFTR deficiency increases total and active RhoA.

Rho GTPases are key regulators of phagocytosis mediated by Fcγ, complement, and efferocytosis receptors (6, 16, 18). Efferocytosis is inhibited by RhoA, whereas Rac-1 and Cdc42 stimulate the process (31, 52). Because total and active RhoA have recently been reported to be increased in CF cell lines and nasal epithelium (28), we wanted to determine whether it was involved in defective efferocytosis by CFTR^−/− epithelium. Total, basal-active, and GTPγS-activated RhoA were consistently increased in IB3-1 epithelial cells compared with their C38 controls (Fig. 5A), whereas only basal-active and GTPγS-activated RhoA were increased in CFTR^−/− primary airway epithelium (Fig. 5B). One of the downstream effects of active RhoA is the assembly of actin stress fibers. Consistent with the finding that active RhoA was elevated in IB3-1 cells, we found that formation of actin stress fibers was enhanced in serum-starved or LPA-treated IB3-1 cells, but not in IB3-1 cells treated with complete medium (Fig. 5C). These data confirm and extend the finding that CFTR deficiency causes an imbalance in RhoA that could be predicted to inhibit efferocytosis (52).

Fig. 5.

CFTR deficiency increases RhoA activity. A: Western blot analysis of total RhoA protein and baseline and GTPγS-loaded RhoA activity in C38 and IB3-1 epithelial cells. Statistical analysis was performed on densitometry readings of total or active RhoA relative to its β-actin control for C38 and IB3–1 cells (n = 3). B: Western blot analysis of total RhoA protein and baseline and GTPγS-loaded RhoA activity in CFTR^+/+ and CFTR^−/− primary airway epithelial cells. Representative blots of 2 replicates are shown for CFTR^+/+ and CFTR^−/− cells. C: actin polymerization (green) and stress fiber formation in C38 and IB3-1 cells exposed to complete medium (media control), serum starvation for 6 h, or lysophosphatidic acid (LPA) for 10 min. Serum starvation and LPA enhanced actin stress fibers (white arrow) more in IB3-1 than in C38 cells. Fluorescence images are representative of 4 independent experiments.

RhoA and Rho kinase inhibitors restore phagocytic competency in CFTR^−/− epithelial cells.

RhoA inhibits efferocytosis through its downstream effector Rho kinase (52). Inhibitors of RhoA (C3 transferase) and Rho kinase (Y-27632) antagonize the effects of RhoA and increase efferocytosis by macrophages (52). Accordingly, C3 transferase completely restored efferocytosis in IB3-1 cells without altering binding (Fig. 6, A and B). Similarly, Y-27632 restored efferocytosis by IB3-1 cells (Fig. 6C) and CFTR^−/− primary airway epithelial cells (Fig. 6E) in a dose-dependent manner but had no effect on binding (Fig. 6, D and F). These results suggest that high RhoA activity in CFTR^−/− cells is directly linked to impaired efferocytosis.

Fig. 6.

Inhibitors of RhoA, Rho kinase, and amiloride-sensitive ion channels restore phagocytic competency in CFTR^−/− cells. CFTR^+/+ and CFTR^−/− airway epithelial cells were treated with or without C3 transferase (A and B), Y-27632 (C–F), or amiloride (G and H) and then cocultured with no cells (NC) or apoptotic Jurkat T cells (ApoJkts). Values are means ± SE for 3–6 replicates per group. C3 transferase, Y-27632, and amiloride corrected defective efferocytosis in untreated IB3-1 and CFTR^−/− primary epithelial cells without affecting binding. *Significantly different from identically treated CFTR^+/+ cells (P < 0.05) **Significantly different from untreated CFTR^−/− cells (P < 0.05). †Significantly different from untreated C38 cells (P < 0.05).

Amiloride restores phagocytic competency in CFTR-deficient cells.

CFTR (1, 33) and the RhoA/Rho kinase pathway (9, 44, 46) regulate several amiloride-sensitive ion channels, including the epithelial Na⁺ channel (ENaC) and Na⁺/H⁺ exchanger-3 (NHE3). These channels also interact directly, or indirectly, with the actin cytoskeleton (7–9), and, interestingly, NHE1, the major epithelial isoform in the lung, appears to directly regulate the actin cytoskeleton, downstream of RhoA (9). These findings suggest that CFTR-regulated, amiloride-sensitive ion channels may also play a role in efferocytosis. Accordingly, the lowest dose of amiloride (0.3 mM) corrected defective efferocytosis by IB3-1 epithelial cells without altering binding (Fig. 6, G and H). In contrast, the highest dose of amiloride (3 mM) decreased efferocytosis in C38 cells, an effect that was expected because of the known inhibitory effect of amiloride on efferocytosis (19). Taken together, these data suggest that dysfunctional CFTR inhibits efferocytosis through a mechanism involving RhoA, Rho kinase, and an amiloride-sensitive ion channel.

Apoptotic cells induce IL-8 production by CFTR^−/− epithelial cells in a Rho kinase-independent manner.

Apoptotic cells normally exert potent anti-inflammatory effects in vitro and in vivo by suppressing a variety of proinflammatory mediators, including IL-8 (54). However, since CFTR deficiency impaired efferocytosis (Figs. 1 and 2), we hypothesized that apoptotic cells would not suppress IL-8 release by CFTR^−/− epithelial cells.

Apoptotic or viable Jurkat T cells were cocultured with C38 and IB3-1 epithelial cell lines in the presence or absence of inflammatory stimuli [TNFα (220 U/ml) and IL-1β (50 ng/ml)]. Apoptotic cells increased IL-8 release by stimulated IB3-1 cells compared with stimulated IB3-1 cells without coculture (Fig. 7A). The effect of apoptotic cells appeared to be independent of cell type, because apoptotic neutrophils also increased IL-8 release by stimulated IB3-1 cells (data not shown). TNFα and IL-1β also increased IL-8 release by IB3-1 cells compared with C38 controls, and this was independent of coculture (Fig. 7A). Therefore, apoptotic cells have a proinflammatory effect on CFTR^−/− IB3-1 epithelial cells.

Fig. 7.

Apoptotic cells enhance IL-8 release from CFTR^−/− epithelial cells in a Rho kinase-independent manner. A: IL-8 release in C38 and IB3-1 epithelial cells cocultured with no cells (NC), viable cells (V), or apoptotic Jurkat T cells (Apo) in the presence of no stimulus (No Stim), TNFα, or IL-1β. Values are means ± SE for 4–6 replicates per group. *Significantly different from IB3-1 cells without coculture (P < 0.05). **Significantly different from identically treated C38 cells (P < 0.05). B: IL-8 release in CFTR^+/+ and CFTR^−/− primary epithelial cells treated as described in A. Values are means ± SE for 5 replicates per group. *Significantly different from CFTR^−/− cells without coculture (P < 0.05). C: test for involvement of the RhoA/Rho kinase pathway in apoptotic cell-enhanced IL-8 release from CFTR^−/− cells. C38 and IB3-1 epithelial cells were cocultured with no cells or apoptotic Jurkat T cells in the presence or absence of IL-1β and in the presence or absence of the Rho kinase inhibitor Y-27632 (1 and 5 μM), and supernatants were collected and analyzed for IL-8. Values are means ± SE for 3 replicates per group. *Significantly different from identically treated C38 cells (P < 0.05). **Significantly different from identically treated IB3-1 cells without coculture (P < 0.05).

These findings were confirmed in CFTR^−/− primary airway epithelial cells and CFTR^+/+ controls in experiments identical to those described in Fig. 7A. Apoptotic cells again enhanced IL-8 release, but by both unstimulated and stimulated CFTR^−/− epithelial cells compared with CFTR^+/+ controls (Fig. 7B). We were surprised to find that apoptotic cells did not induce TGFβ1 in CFTR^+/+ or CFTR^−/− epithelial cells (data not shown). Stimulation did not enhance IL-8 release by CFTR^−/− epithelial cells in the presence or absence of cocultured viable Jurkat T cells (Fig. 7B).

Finally, similar experiments were performed to determine whether apoptotic cell-enhanced IL-8 release was dependent on the RhoA/Rho kinase pathway (Fig. 7C). These experiments were performed as described in Fig. 7A, but in the presence or absence of the Rho kinase inhibitor Y-27632. As shown in Fig. 7A, apoptotic cells enhanced IL-8 release by IB3-1 cells, but this effect was not abrogated by inhibition of Rho kinase. Taken together, these data indicate that apoptotic cells enhance, rather than suppress, IL-8 release by CFTR^−/− epithelial cells in vitro in a Rho kinase-independent manner, suggesting an additional mechanism for increased IL-8 in CF.

DISCUSSION

Because a large reservoir of dead (apoptotic and necrotic) cells is present in the CF airway (48, 53), we sought to determine whether CFTR deficiency specifically interferes with epithelial efferocytosis and its normal suppression of inflammatory mediator release. Here we show that defective CFTR directly impedes epithelial cell efferocytosis and that this defect in phagocytosis extends to ingestion by the Fcγ receptor. The mechanism of this CFTR-dependent effect is related to dysregulated Rho GTPases, because 1) total and active RhoA are increased in CFTR^−/− epithelial cells and 2) inhibitors of RhoA and its downstream effector Rho kinase normalize efferocytosis. RhoA/Rho kinase may be acting downstream through ENaC or NHE1, because amiloride also normalizes defective efferocytosis. The consequences of apoptotic cell accumulation appear to be proinflammatory, because apoptotic cells enhance IL-8 release by CFTR-deficient epithelial cells.

Epithelial cells are capable of ingesting apoptotic cells of virtually the same size in vitro and in vivo (20, 29, 37, 57). Most cell types, including mesenchymal cells, macrophages, fibroblasts, mesangial cells, hepatocytes, glia, and epithelial cells, have the capacity to ingest apoptotic cells (18, 37, 57, 60). One of the best examples of this may be the “macrophageless” PU.1 knockout mouse (60). These mice have no mature macrophages, yet when the interdigital web undergoes massive apoptosis during normal intrauterine development, neighboring mesenchymal cells “stand in” for the macrophages and ingest the apoptotic cells, albeit at one-third of the rate of ingestion by macrophages.

CFTR deficiency impaired the ability of airway epithelium to remove apoptotic cells, which may be related to its membership in the ABC transporter superfamily. Similar to CFTR, several ABC transporters, including ced-7 (61), ABC-A1 (32), and ABC-A7 (21), have been implicated in efferocytosis. In Caenorhabditis elegans, two signaling pathways govern corpse removal; both converge at a common effector, ced-10 (Rac-1) (27). The first pathway includes ced-2 (CrkII), ced-5 (Dock180), and ced-12 (ELMO), and the second includes ced-1 (LRP), ced-6 (GULP), and ced-7 (ABC-A1 or ABC-A7) (27). The exact mechanism by which ced-7 contributes to the clearance process has not been elucidated, but its presence is required on the phagocyte and apoptotic cell for maximum corpse engulfment (61). Similarly, one putative ced-7 mammalian homolog, ABC-A1 (61), which causes Tangier disease in humans, has been implicated as a regulator of efferocytosis in mice; it also appears to be required on the phagocyte and apoptotic cell, perhaps because of its ability to affect redistribution of membrane PS (18). In addition, similar to CFTR, ABC-A1 is also a cAMP-dependent, glibenclamide-sensitive anion transporter. Our knowledge about ABC-A7 is evolving, but during macrophage efferocytosis, ABC-A7 concentrates to the phagocytic cup, where it colocalizes with LRP and facilitates efferocytosis through LRP-dependent phosphorylation of extracellular signal-related kinase (21).

Dysfunctional CFTR was associated with increased total and active RhoA, which negatively impacted efferocytosis in a Rho kinase-dependent manner. Rho GTPases (e.g., RhoA, Rac-1, and Cdc42) are molecular switches, cycling between off (GDP-bound) and on (GTP-bound) configurations (59). Rho GTPase activity is tightly regulated in three major ways: 1) GDP/GTP binding, 2) lipid modification and subcellular localization, and 3) gene expression (59). RhoA and Rac-1 can also downregulate each other through cross talk (5). The mechanism(s) by which CFTR deficiency enhances total and active RhoA has not been defined, but dysfunctional CFTR may be exerting its effect, at least through altered gene expression and GDP/GTP binding. A subfamily of RhoA-specific guanine nucleotide exchange factors, such as Lbc, Lfc, Lsc, P115RhoGEF, leukemia-associated RhoGEF (LARG), and XPLN, is beginning to emerge; these factors catalyze the conversion of inactive GDP-bound RhoA to its active GTP-bound form and, thereby, could contribute to increased RhoA activity in CFTR^−/− epithelial cells. CFTR deficiency also increases oxidative stress in vitro and in vivo, likely through its role as a glutathione transporter and regulator of NADPH oxidase activity. This mechanism may be particularly important, because oxidative stress activates RhoA and inhibits efferocytosis (35), implying that a similar effect may occur in the CF airway.

Amiloride-sensitive ion channels also appear to be involved with impaired efferocytosis. This is intriguing, because CFTR (1, 33) and RhoA (9, 44, 46) regulate several amiloride-sensitive channels. We speculate that ENaC and/or NHE1 may be involved, because they are present in airway epithelium. ENaC is a particularly attractive candidate, because dysfunctional CFTR increases ENaC activity in airway epithelial cells (33) and lung-specific ENaC overexpression results in a pulmonary phenotype virtually identical to CF (34). In contrast, the ability of CFTR to regulate airway epithelial Na⁺/H⁺ exchange is controversial (40).

How else might CFTR exert its regulatory effect on efferocytosis? CFTR is a transmembrane protein present in the plasma membrane, the trans-Golgi network, recycling endosomes, and the phagocytic cup (3, 11). In the plasma membrane, CFTR exists as part of a macromolecular complex, which is facilitated by protein-protein interactions based on the presence of a PDZ binding motif at the COOH terminus of CFTR. Scaffolding and regulatory proteins containing the type I PDZ domain binding sites, such as CAP70 and the Na⁺/H⁺ exchanger regulatory factors 1 and 2, bind to CFTR at its COOH terminus. These protein-protein associations function as CFTR membrane retention signals (39), regulate CFTR channel activity through formation of CFTR multimers (58), or link CFTR to other ion channels (e.g., NHE3) (1), protein kinase A (45), ezrin-radixin-moesin (4), and the actin cytoskeleton (4). Through these direct and indirect mechanisms, CFTR may have unforeseen regulatory effects on the actin cytoskeleton that may be independent of its ion channel activity. Data presented here suggest that CFTR regulates epithelial cell efferocytosis in a channel-independent manner, perhaps through one of these associations.

Dysfunctional CFTR also suppressed Fcγ receptor-mediated phagocytosis. Similar to efferocytosis, Rac-1 and Cdc42 signaling are required for Fcγ receptor-mediated phagocytosis. In contrast, a role for RhoA is still controversial, because inhibitors of RhoA have been shown to suppress Fcγ receptor-mediated phagocytosis (16) or to have no effect (6). In these studies, inhibitors or dominant-negative constructs were used to probe the RhoA pathway; no studies have examined the effect of endogenous RhoA activation, constitutively active constructs, or overexpression. It is interesting that RhoA accumulates at the phagocytic cup during Fcγ receptor-mediated phagocytosis (6), suggesting that it may influence the process. Because RhoA indirectly antagonizes Rac-1 signaling (5), it remains possible that enhanced RhoA activity in CFTR^−/− cells may indirectly suppress Fcγ receptor-mediated phagocytosis and, for that matter, efferocytosis.

Apoptotic cells induced IL-8 release by CFTR^−/− cells, independent of the cell type and in a Rho kinase-independent manner; the magnitude of this effect is surprising considering that apoptotic cells normally suppress inflammatory mediator release via PS-dependent induction of TGFβ1 (18). Instead, apoptotic cells failed to induce TGFβ1 by CFTR^+/+ or CFTR^−/− epithelial cells. Ineffective efferocytosis might also be expected to cause inflammatory mediator release because of the development of postapoptotic necrosis. Work is underway to further elucidate the mechanism(s) underlying the proinflammatory effect of apoptotic cells. Several possibilities exist. 1) Increased RhoA has been linked to NFκB activation (50) and may play a role in enhanced IL-8 release through one of its downstream effectors. Rhotekin is an attractive candidate, because it can activate NFκB downstream of RhoA. 2) Despite our inability to demonstrate the presence of LRP on C38 or IB3-1 cells, apoptotic cell interaction with CFTR^−/− cells may occur through an LRP-dependent, rather than a PS recognition-dependent, mechanism. Because LRP engagement has been shown to activate NFκB and induce inflammatory mediator release, efferocytosis, largely through an LRP-dependent mechanism, may enhance proinflammatory mediator release. 3) TGFβ1 signaling is defective in cells expressing dysfunctional CFTR (24), suggesting that were TGFβ1 to have been induced, it may not have suppressed IL-8. We were not able to demonstrate elevated TGFβ1 in the supernatants of C38 or IB3-1 cells in response to coculture with apoptotic cells; however, changes in TGFβ1 may be difficult to detect by ELISA.

As intriguing as these results are, we present them with an important caveat: the importance of lung epithelium in the removal of apoptotic cells in vivo has not yet been demonstrated. This is unlike other settings, such as the involuting mammary gland (37), the retinal pigment epithelium (14, 15, 62), or injured kidney tubules (20), where epithelial cells have been shown to have a definitive role in the clearance of apoptotic cells. We know that lung epithelium participates in the removal of apoptotic cells during lung development (29). In the developed lung, many questions remain, not the least of which are as follows. 1) Under what conditions (basal or inflammatory) might the lung epithelium become phagocytic? 2) What are the consequences of this phagocytic activity, and which epithelial cells participate? 3) Are all apoptotic targets recognized equally, or is there a hierarchy? 4) Are transmigrating or neighboring apoptotic cells recognized by lung epithelium more effectively than cells within the lumen?

Ultimately, ineffective efferocytosis and accumulation of apoptotic cells in CF (2, 48, 53) may present fertile ground for novel therapies, especially if efferocytosis is defective in macrophages (53) and epithelial cells. Drugs that specifically block the RhoA/Rho kinase pathway may, if applied early, control the apoptotic cell burden and maintain lung homeostasis. Statins block RhoA, enhance efferocytosis (38), and are increasingly recognized to possess broad anti-inflammatory effects that could be immediately applied to CF (54). In addition, peroxisome proliferator-activated receptor-γ agonists, macrolide antibiotics, and glucocorticoids also promote efferocytosis and suppress inflammation, and some of these have a recognized role in the treatment of CF lung inflammation (54). Inhaled amiloride has been disappointing, in that it does not demonstrate clinical efficacy; perhaps earlier intervention and better pharmacological profiles would produce a different outcome by more effectively maintaining lung quiescence before other mechanisms take hold.

GRANTS

This work was support by National Institutes of Health Grants HL-072018 and HL-088138 (to R. W. Vandivier), GM-61031, HL-68864, and HL-088138 (to P. M. Henson), and P50 DK-49096-06 and HL-090669 (to G. P. Downey) and the Harold and Mary Zirin Chair in Pulmonary Cell Biology of National Jewish Health (to G. P. Downey). R. W. Vandivier was also supported by a CIA Award from the Flight Attendant Medical Research Institute (072001-CIA).