Airway epithelial cells have a major role in initiating inflammation in response to bacterial pathogens. Through the immediate induction of CXCL8 and cytokine expression, polymorphonuclear cells are mobilized and activated to eradicate the infecting organisms. However, the influx of polymorphonuclear cells and the effects of their toxic exoproducts impede respiratory function. We postulated that respiratory epithelial cells must also participate in the regulation of their own proinflammatory signaling. Both Staphylococcus aureus and Pseudomonas aeruginosa were found to potently activate IL-6 expression immediately upon contact with epithelial cells, and by 1 h induced TNF-α converting enzyme (TACE) transcription. By 4 h of bacterial exposure, TACE colocalized with IL-6Rα on the apical surface of airway cells, and by 24 h, soluble IL-6Rα accumulated in the cell culture supernatant. Epithelial IL-6 and soluble IL-6Rα were shown to participate in trans-signaling, interacting with membrane-associated gp130 to activate CCL-2 expression and inhibit additional CXCL8 production. Thus, bacteria are physiological activators of TACE expression, which provides a mechanism to regulate inflammatory signaling that is initiated by airway epithelial cells.

The active participation of airway epithelial cells in proinflammatory signaling is well recognized. Airway cells produce the polymorphonuclear cell (PMN) 4 chemokine CXCL8, as well as the cytokines G-CSF, GM-CSF, and IL-6, which contribute to the recruitment, activation, and persistence of PMNs in response to the presence of bacterial pathogens in the airway lumen (1, 2, 3). Because the accumulation of PMNs in the airway and the release of their potentially damaging products, such as reactive oxygen intermediates and elastase, are not without harm to the host, numerous regulatory mechanisms control PMN-dominated inflammation. Although there are several anti-inflammatory cytokines that modulate proinflammatory signaling at mucosal surfaces, such as IL-10 and TGF-β (4, 5, 6), the sources of these cytokines are not clearly defined.

Because mucosal epithelial cells have numerous mechanisms to activate proinflammatory signaling, they should also have corresponding signaling pathways that regulate this response. One of the cytokines produced by mucosal cells, IL-6, has both pro- and anti-inflammatory effects (7, 8, 9). Epithelial responsiveness to IL-6 is dependent upon the presence of two potential receptors, the ubiquitously expressed gp130 and IL-6Rα (gp80), which is more limited in its distribution (8, 10). A soluble form of the IL-6Rα (sIL-6Rα) can be released from cell membranes through the activity of proteases, including TNF-α converting enzyme (TACE), a disintegrin and metalloprotease (ADAM 17) (11) and aminopeptidase regulator of TNFR1 shedding (ARTS-1) (12). The regulation of these key enzymes is not well understood, nor are the physiological signals for their expression and mobilization to the cell surface defined (13). Production of sIL-6Rα can also be the result of a differential mRNA splicing (DS sIL-6R) (11). Soluble IL-6Rα binds to IL-6, forming a ligand-receptor complex that interacts with gp130 in a high affinity interaction termed trans-signaling. This interaction initiates CCL-2 expression, which heralds the shift from acute PMN-dominated inflammation to macrophage/monocyte recruitment and contributes to the clearance of apoptotic PMNs (7, 14).

PMNs and macrophages have been considered to be the major source of shed sIL-6Rα (15, 16). In the airway lumen, sIL-6Rα could be produced by either immune cells or the epithelium. If airway epithelial cells were to express IL-6Rα, its cleavage from the cell surface and secretion could provide a mechanism to regulate inflammation through IL-6/sIL-6Rα/gp130 signaling, which, in turn, would drive epithelial CCL-2 production. In the studies presented in this report, we demonstrate that both Gram-positive and Gram-negative pulmonary pathogens directly activate the regulatory components of the IL-6 signaling cascade in airway epithelial cells. Airway cells produce IL-6 and have a major role in regulating inflammation through activation of TACE, IL-6Rα shedding, and CCL-2 expression.

16HBE and 1HAEo- cells, human airway epithelial cell lines (D. Gruenert, California Pacific Medical Center Research Institute, San Francisco, CA) were grown as previously detailed (2, 17). Primary airway epithelial cells isolated from human nasal polyps were grown on Transwell-Clear filters (Corning Costar) in M3 medium as previously described (18). For heat-killed Staphylococcus aureus and Pseudomonas aeruginosa preparations, bacteria were grown in CyGP or Luria-Bertoni medium overnight, then resuspended in MEM (Invitrogen Life Technologies) and heated for 60 min at 60°C. S. aureus-purified protein A and P. aeruginosa LPS were obtained from Sigma-Aldrich. Peptidoglycan was obtained from Calbiochem. P. aeruginosa flagella was prepared as previously described (19). The mAbs used were anti-gp80 (IL-6Rα; BioSource International) and anti-gp130 (BD Biosciences). Goat polyclonal anti-TACE Ab was obtained from Santa Cruz Biotechnology.

16HBE cells, weaned from serum for 24 h, were exposed to heat-killed S. aureus or P. aeruginosa (108 CFU) for 1, 4, or 24 h. The effect of MAPK inhibitors was tested by pretreating the cells for 60 min with 6 μM SB202190 (Calbiochem) or 10 μM UO126 (Calbiochem). For depletion of Ca2+, BAPTA (6 μM; Molecular Probes) was used. IL-6 in the supernatant was measured by ELISA as previously described (2).

16HBE cells grown in 12-well plates to 50–70% confluence were transiently transfected using FuGene 6.0 (Roche) with pNF-κB-luciferase (0,5 μg/ml), pAP-1-luciferase (0.5 μg/ml), or pc/EBP-luciferase (0.5 μg/ml) (Stratagene). After 16 h, cells were weaned from serum for 24 h and stimulated with S. aureus or P. aeruginosa for 6 h. Luciferase assays were performed as previously described (2).

16HBE were grown to confluence on Transwell-Clear filters (Corning-Costar) with an air-liquid interface to form polarized monolayers. Cells were fixed with 4% paraformaldehyde for 20 min at room temperature and incubated with 5% normal serum blocking solution for 30 min at room temperature. Primary Abs were added for 1 h at room temperature, followed by three 5-min washes. Alexa Fluor 488- and 594-conjugated secondary Abs (Molecular Probes) were added for 1 h and washed three times. Filters were removed from Transwells using a scalpel and were mounted with Vectashield (Vector Laboratories) onto glass slides.

16HBE cells were washed three times after stimulation and incubated with 5% normal serum for 30 min at room temperature. Primary Abs were added for 1 h at room temperature, followed by three washes. Alexa Fluor 488-conjugated secondary Ab (Molecular Probes) was added for 1 h at room temperature. Cells were then washed, fixed in 1% paraformaldehyde, and analyzed with a FACSCalibur using CellQuest software (BD Biosciences).

16HBE cells, weaned from serum for 24 h, were exposed to heat-killed S. aureus or P. aeruginosa (108 CFU) for 1, 4, or 24 h or to protein A (200 μg/ml), peptidoglycan (10 μg/ml), or flagella (10 μg/ml) for 24 h. Soluble IL-6Rα and CCL-2 in the supernatants were detected by using DuoSet ELISA for human sIL-6Rα and human CCL-2, respectively (R&D Systems). For TACE inhibition experiments, cells were preincubated with TNF-α protease inhibitor-1 (TAPI-1) (Calbiochem) for 30 min, then stimulated in the presence of TAPI-1. Soluble gp130 (Calbiochem) was added for 30 min before and during stimulation.

Epithelial cells were grown in six-well plates to 80% confluence and were weaned from serum overnight. After stimulation with heat-killed bacteria, flagella, or medium alone, cells were lysed, and RNA was isolated using the Qiagen RNeasy Mini Kit. cDNA was made from 1 μg of RNA using an iScript synthesis kit (Bio-Rad). For IL-6R RT-PCR, Taq polymerase (Roche) was used, and 25 or 35 cycles were run with denaturation at 95°C for 1 min, annealing at 60°C for 40 s, and extension at 72°C for 40 s. The primers used were 5′-CAGCTGAGAACGAGGTGTCC-3′ and 5′-GCAGCTTCCACGTCTTCTTGA-3′. For quantitative real-time PCR of TACE and CXCL8, cDNA amplification was performed in a Light Cycler using the DNA Master SYBR Green I kit (Roche) according to the manufacturer’s instructions. Primers used for TACE amplification were 5′-ACCTGAAGAGCTTGTTCATCGAG-3′ and 5′-CCATGAAGTGTTCCGATAGATGTC-3′, and 35 cycles were run with denaturation at 95°C for 8 s, amplification at 54°C for 15 s, and extension at 72°C for 12 s. The primers used for CXCL8 amplification were 5′-TACTCCAAACCTTTCCAACCC-3′ and 5′-AACTTCTCCACAACCCTCTG-3′, and 35 cycles were run with denaturation at 95°C for 8 s, amplification at 55°C for 10 s, and extension at 72°C for 12 s. Amplification of actin was performed on each sample and used as a control for standardization. The primers used for actin amplification were 5′-GTGGGGCGCCCCAGGCACCA-3′ and 5′-CGGTTGGCCTTGGGGTTCAGGGGGG-3′, and 35 cycles were run with denaturation at 95°C for 8 s, amplification at 63°C for 10 s, and extension at 72°C for 12 s.

Two pairs of oligonucleotides containing 19 bp of human TACE were generated as follows: pair 1, 5′-gatccccGTAAGGCCCAGGAGTGTTTttcaagagaAAACACTCCTGGGCCTTACttttggaaa-3′ and 5′-agcttttccaaaaGTAAGGCCCAGGAGTGTTTtctcttgaaAAACACTCCTGGGCCTTAC ggg-3′; and pair 2, 5′-gatccccCATAGAGCCACTTTGGAGAttcaagagaTCTCCAAAGTGGCTCTATG ttttggaaa-3′ and 5′-agcttttccaaaaCATAGAGCCACTTTGGAGAtctcttgaaTCTCCAAAGTGGCTC TATGggg-3′. To construct pRS-TACE-1 and pRS-TACE-2, the oligos were annealed and ligated into BglII and HindIII sites of pRetroSuper vector (pRS) (20). Construct integrity was confirmed by direct sequencing of the plasmid. Packaging of retroviral constructs was conducted in HEK 293T cells (21). 16HBE cells were infected for 18 h in the presence of 4 mg/ml polybrene (Sigma-Aldrich). The pBabe-puro-enhanced GFP was used to monitor the efficiency of transfection to 293T cells and infection. A pRS-scramble plasmid (pRS-sc) was used as a control by cloning the sequence ggcagttccaccccagtgc into pRS as described for pRS-TACE.

S. aureus and P. aeruginosa stimulated the production of biologically significant amounts of IL-6 in 16HBE airway epithelial cells (p < 0.01; Fig. 1, A and B). Purified bacterial components, including staphylococcal protein A and peptidoglycan, as well as Pseudomonas flagella also induced IL-6 production by epithelial cells in a dose-dependent fashion (Fig. 1, C–E). As shown for other proinflammatory cytokines, epithelial IL-6 expression involves the generation of Ca2+ fluxes, MAPKs, and the expected transcription factors. IL-6 production induced by S. aureus, P. aeruginosa, as well as purified bacterial components was significantly inhibited (p < 0.05 and p < 0.01) in the presence of p-38 and ERK1/2 MAPK inhibitors (Fig. 2,A) as well as BAPTA (p < 0.01; Fig. 2,B). The transcription factors, AP-1, c/EBP, and NF-κB, which have been shown to be involved in IL-6 production (22, 23) in other cell types, were also activated in response to S. aureus and P. aeruginosa, as shown by luciferase reporter assays (p < 0.01; Fig. 2 C).

FIGURE 1.

Epithelial IL-6 production in response to bacteria. AE, IL-6 was assayed by ELISA after exposure of 16HBE cells to medium alone (C, control), heat-killed S. aureus, heat-killed P. aeruginosa, or the indicated amounts of purified bacterial products. PGN, peptidoglycan. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.01; ∗∗, p < 0.05.

FIGURE 1.

Epithelial IL-6 production in response to bacteria. AE, IL-6 was assayed by ELISA after exposure of 16HBE cells to medium alone (C, control), heat-killed S. aureus, heat-killed P. aeruginosa, or the indicated amounts of purified bacterial products. PGN, peptidoglycan. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.01; ∗∗, p < 0.05.

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FIGURE 2.

Signaling pathways involved in IL-6 production by airway epithelial cells. 16HBE cells were pretreated with the p38 MAPK inhibitor SB202190, the MEK1/2 inhibitor UO126 (A), or BAPTA (B) and stimulated with bacteria or purified bacterial products as indicated. Culture supernatants were assayed for IL-6 by ELISA. PAO1, P. aeruginosa; PGN, peptidoglycan. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.01; ∗∗, p < 0.05. C, 16HBE cells were transiently transfected with NF-κB, AP-1, or c/EBP luciferase reporter constructs. After stimulation, luciferase activity was measured and compared with unstimulated control cells. ∗, p < 0.01.

FIGURE 2.

Signaling pathways involved in IL-6 production by airway epithelial cells. 16HBE cells were pretreated with the p38 MAPK inhibitor SB202190, the MEK1/2 inhibitor UO126 (A), or BAPTA (B) and stimulated with bacteria or purified bacterial products as indicated. Culture supernatants were assayed for IL-6 by ELISA. PAO1, P. aeruginosa; PGN, peptidoglycan. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.01; ∗∗, p < 0.05. C, 16HBE cells were transiently transfected with NF-κB, AP-1, or c/EBP luciferase reporter constructs. After stimulation, luciferase activity was measured and compared with unstimulated control cells. ∗, p < 0.01.

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To respond to IL-6 signaling, epithelial cells must express the appropriate IL-6Rs. We found both the ubiquitous receptor gp130 as well as IL-6Rα (gp80) on the surface of 16HBE cells (Fig. 3,A). By confocal imaging, gp80 appeared to be distributed chiefly on the apical surface, whereas gp130 had both apical and basolateral distributions (Fig. 3,B). These airway cells also expressed DS sIL-6R, as shown by RT-PCR (Fig. 3,C). To establish whether airway cells, like leukocytes, actively shed a soluble form of the IL-6Rα we performed a kinetic study after the release of IL-6Rα from the surface of 16HBE cells in response to either S. aureus or P. aeruginosa (Fig. 4,A). In contrast to the relatively rapid activation of IL-6 secretion, significant amounts of sIL-6Rα were not found in the epithelial culture supernatant until 24 h of S. aureus exposure. The continued presence of the bacteria was necessary for shedding to occur, because culture supernatants harvested 23 h after a 1-h exposure to bacteria did not contain sIL-6Rα. Similar levels of IL-6Rα shedding were observed using human airway cells in primary culture (Fig. 4,B). P. aeruginosa also stimulated IL-6Rα shedding, but, due to destruction of the monolayers after 4 h of exposure to even heat-killed organisms, longer time points could not be studied. Epithelial stimulation for 24 h with purified staphylococcal protein A or P. aeruginosa flagella induced significant amounts of IL-6Rα shedding (Fig. 4,C). A concomitant decrease in surface IL-6Rα on the intact epithelial monolayers correlated with the appearance of shed receptor in the culture supernatant (Fig. 4,D). The sIL-6R detected was not a consequence of increased expression of DS sIL-6R, as determined by RT-PCR (Fig. 4,E). Because S. aureus and P. aeruginosa induce CXCL8 as well as IL-6 (2), and CXCL8 can induce IL-6Rα shedding in neutrophils (15), we determined whether either of these cytokines was responsible for IL-6Rα shedding by epithelial cells (Fig. 4,F). Stimulation of epithelial cells with IL-6 or CXCL8 did not induce IL-6Rα shedding, nor did conditions known to induce high levels of CXCL8 (supernatants collected 23 h after 1-h exposure to S. aureus; Fig. 4 A).

FIGURE 3.

IL-6R complex expression on airway epithelial cells. A, 16HBE cells under basal conditions were stained for surface expression of gp80 (IL-6Rα) and gp130. The shadow histogram is the isotype control; the black line indicates the specific Ab. B, Confocal images (Z sections) of polarized 16HBE stained for gp80 and gp130 are shown. C, RT-PCR for membrane-bound IL-6Rα (219-bp band) and DS sIL-6Rα (125-bp band). Lanes 13, 16HBE, 1HAEo- (human airway epithelial), and THP-1 cells (positive control), respectively.

FIGURE 3.

IL-6R complex expression on airway epithelial cells. A, 16HBE cells under basal conditions were stained for surface expression of gp80 (IL-6Rα) and gp130. The shadow histogram is the isotype control; the black line indicates the specific Ab. B, Confocal images (Z sections) of polarized 16HBE stained for gp80 and gp130 are shown. C, RT-PCR for membrane-bound IL-6Rα (219-bp band) and DS sIL-6Rα (125-bp band). Lanes 13, 16HBE, 1HAEo- (human airway epithelial), and THP-1 cells (positive control), respectively.

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FIGURE 4.

IL-6Rα shedding by airway epithelial cells. 16HBE cells (A and C) or human primary epithelial cells (B) were exposed to medium alone (C, control), heat-killed S. aureus, or heat-killed P. aeruginosa (PAO1) for the times indicated or to purified bacterial components for 24 h. Soluble IL-6Rα was assessed in the supernatant by ELISA. Data represent the mean and SD from sextuplicate wells (A and C). ∗, p < 0.01. D, 16HBE cells were stimulated with heat-killed S. aureus or P. aeruginosa (PAO1) for the times indicated, and IL-6Rα on the surface was quantified by flow cytometry. Data represent the relative mean fluorescence intensity (MFI) compared with unstimulated cells (C, control). E, RT-PCR for IL-6Rα and DS sIL-6R was performed on RNA from 16HBE cells untreated or stimulated with heat-killed S. aureus or P. aeruginosa for the times indicated. RT-PCR for β-actin is shown as the control. F, 16HBE cells were stimulated with the indicated amounts of IL-6 or CXCL8 for 24 h. Soluble IL-6Rα in the supernatant was assessed by ELISA. Data represent the mean and SD from sextuplicate wells.

FIGURE 4.

IL-6Rα shedding by airway epithelial cells. 16HBE cells (A and C) or human primary epithelial cells (B) were exposed to medium alone (C, control), heat-killed S. aureus, or heat-killed P. aeruginosa (PAO1) for the times indicated or to purified bacterial components for 24 h. Soluble IL-6Rα was assessed in the supernatant by ELISA. Data represent the mean and SD from sextuplicate wells (A and C). ∗, p < 0.01. D, 16HBE cells were stimulated with heat-killed S. aureus or P. aeruginosa (PAO1) for the times indicated, and IL-6Rα on the surface was quantified by flow cytometry. Data represent the relative mean fluorescence intensity (MFI) compared with unstimulated cells (C, control). E, RT-PCR for IL-6Rα and DS sIL-6R was performed on RNA from 16HBE cells untreated or stimulated with heat-killed S. aureus or P. aeruginosa for the times indicated. RT-PCR for β-actin is shown as the control. F, 16HBE cells were stimulated with the indicated amounts of IL-6 or CXCL8 for 24 h. Soluble IL-6Rα in the supernatant was assessed by ELISA. Data represent the mean and SD from sextuplicate wells.

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TACE, a metalloprotease of the ADAM family, is responsible for IL-6Rα shedding in immune cells and is known to be expressed in pulmonary cells (11, 13). In response to S. aureus or P. aeruginosa, TACE mRNA expression was significantly up-regulated as early as 1 h after stimulation (Fig. 5,A). Stimulation of epithelial cells with flagella for 4 h induced a significant increase in TACE mRNA (Fig. 5,A). ARTS-1 has also been shown to participate in IL-6Rα shedding (12). Bacterial stimulation of airway epithelial cells induced a significant increase in ARTS-1 mRNA expression (Fig. 5,B). Flagella-induced IL-6Rα shedding seemed to be independent from ARTS-1, because no transcriptional activation of this aminopeptidase was observed (Fig. 5,B). Although TACE was expressed under basal conditions on the surface of epithelial cells, there was an increase in surface expression after exposure to S. aureus, P. aeruginosa, or flagella (Fig. 5,C). TACE colocalized with the IL-6Rα at the apical surface of polarized airway cells after bacterial and flagella stimulations (Fig. 5 D).

FIGURE 5.

TACE activation by bacteria. TACE mRNA levels (A), ARTS-1 mRNA levels (B), and TACE surface expression (C) were analyzed in 16HBE cells under unstimulated conditions (C, control) or after stimulation with heat-killed S. aureus or P. aeruginosa (PAO1) or flagella. Values for real-time PCR were normalized to β-actin and are shown as the fold change in expression relative to the endogenous level in unstimulated cells. Each bar represents the mean of triplicate samples. ∗, p < 0.01; ∗∗, p < 0.001. D, Polarized 16HBE cells grown in Transwells, stimulated with heat-killed S. aureus or P. aeruginosa or flagella, were stained for TACE (green) and IL-6Rα (red) and analyzed by confocal imaging. Colocalization of TACE and IL-6Rα appears yellow.

FIGURE 5.

TACE activation by bacteria. TACE mRNA levels (A), ARTS-1 mRNA levels (B), and TACE surface expression (C) were analyzed in 16HBE cells under unstimulated conditions (C, control) or after stimulation with heat-killed S. aureus or P. aeruginosa (PAO1) or flagella. Values for real-time PCR were normalized to β-actin and are shown as the fold change in expression relative to the endogenous level in unstimulated cells. Each bar represents the mean of triplicate samples. ∗, p < 0.01; ∗∗, p < 0.001. D, Polarized 16HBE cells grown in Transwells, stimulated with heat-killed S. aureus or P. aeruginosa or flagella, were stained for TACE (green) and IL-6Rα (red) and analyzed by confocal imaging. Colocalization of TACE and IL-6Rα appears yellow.

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To document that TACE functions to cleave the IL-6Rα, we quantified S. aureus-induced IL-6Rα shedding in the presence of TAPI-1, a TACE inhibitor, demonstrating a dose-dependent effect (p < 0.001; Fig. 6,A). Because TAPI-1 may have an inhibitory effect on some metalloproteases other than TACE, we also confirmed that TACE is responsible for IL-6Rα shedding in epithelial cells by using RNA interference. Depletion of TACE significantly reduced endogenous as well as S. aureus- and flagella-induced IL-6Rα shedding (Fig. 6,B and C, pRS-TACE-1). IL-6Rα shedding correlated with the amount of TACE present in the epithelial cells, as shown by comparing IL-6Rα shedding in cells expressing pRS-TACE-1 or pRS-TACE-2 (Fig. 6,B) and demonstrating the effects on protein expression by immunoblot (Fig. 6,E). The effect of TACE depletion on TNFR1 shedding is shown as a control (Fig. 6 D).

FIGURE 6.

Bacteria-induced IL-6Rα shedding is TACE dependent. A, 16HBE cells were stimulated with heat-killed S. aureus in the presence or the absence of increasing amounts of TAPI-1, and sIL-Rα shed into the medium was analyzed by ELISA. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.001. BD, 16HBE cells expressing pRS-TACE (pRS-T) or pRS-sc were stimulated with S. aureus or flagella, and shed sIL-6Rα (B and C) or sTNFR1 (D) was determined by ELISA. ∗, p < 0.01. E, Whole cell lysates from 16HBE cells expressing pRS-TACE or pRS-sc were immunoblotted for TACE and actin. The adjusted volume for each band was determined by densitometry and standardized by actin expression.

FIGURE 6.

Bacteria-induced IL-6Rα shedding is TACE dependent. A, 16HBE cells were stimulated with heat-killed S. aureus in the presence or the absence of increasing amounts of TAPI-1, and sIL-Rα shed into the medium was analyzed by ELISA. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.001. BD, 16HBE cells expressing pRS-TACE (pRS-T) or pRS-sc were stimulated with S. aureus or flagella, and shed sIL-6Rα (B and C) or sTNFR1 (D) was determined by ELISA. ∗, p < 0.01. E, Whole cell lysates from 16HBE cells expressing pRS-TACE or pRS-sc were immunoblotted for TACE and actin. The adjusted volume for each band was determined by densitometry and standardized by actin expression.

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Trans-signaling, the interaction of IL-6, soluble IL-6Rα, and cell-associated gp130, initiates CCL-2 production in leukocytes (7). To determine whether a similar pathway is activated in airway epithelial cells, we first established that bacterial stimulation induces CCL-2 production (Fig. 7, A and B). A small increase in CCL-2 was observed 4 h after S. aureus stimulation, and significantly greater amounts were detected after 24 h of stimulation (Fig. 7,A), when both IL-6 (Fig. 1,A) and shed sIL-6Rα (Fig. 4,A) were present in the extracellular medium. These levels of CCL-2 were not achieved by 24 h after 1-h exposure to S. aureus (Fig. 7,A). Exposure to P. aeruginosa far in induced CCL-2 production. Twenty-four-hour stimulation with P. aeruginosa flagella induced significant amounts of CCL-2 (Fig. 7,B). CCL-2 production was not observed in cells lacking TACE expression (Fig. 7 C), which is consistent with a requirement for IL-6Rα shedding and trans-signaling to promote CCL-2 production in epithelial cells.

FIGURE 7.

CCL-2 production in epithelial cells. A and B, 16HBE cells were exposed to medium alone (C, control), heat-killed S. aureus, heat-killed P. aeruginosa (PAO1), or flagella. CCL-2 production was assessed in the supernatant by ELISA. Data represent the mean and SD from sextuplicate wells (A). ∗, p < 0.01 (compared with control); a, p < 0.01 (compared with 4 h). C, 16HBE cells expressing pRS-TACE (pRS-T-1) or pRS-sc were stimulated with flagella, and CCL-2 production was determined by ELISA. ∗, p < 0.01. D, 16HBE cells were stimulated with heat-killed S. aureus in the presence or the absence of increasing amounts of sgp130, and CCL-2 production was determined by ELISA. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.01.

FIGURE 7.

CCL-2 production in epithelial cells. A and B, 16HBE cells were exposed to medium alone (C, control), heat-killed S. aureus, heat-killed P. aeruginosa (PAO1), or flagella. CCL-2 production was assessed in the supernatant by ELISA. Data represent the mean and SD from sextuplicate wells (A). ∗, p < 0.01 (compared with control); a, p < 0.01 (compared with 4 h). C, 16HBE cells expressing pRS-TACE (pRS-T-1) or pRS-sc were stimulated with flagella, and CCL-2 production was determined by ELISA. ∗, p < 0.01. D, 16HBE cells were stimulated with heat-killed S. aureus in the presence or the absence of increasing amounts of sgp130, and CCL-2 production was determined by ELISA. Data represent the mean and SD from sextuplicate wells. ∗, p < 0.01.

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A soluble form of gp130 (sgp130) is the natural inhibitor of sIL-6Rα trans-signaling responses. Soluble gp130 binds to the IL-6/sIL-6Rα complex without affecting the interaction of IL-6 with membrane IL-6Rα (24). Thus, by using purified sgp130, we confirmed that epithelial CCL-2 production in response to bacteria was due to trans-signaling. CCL-2 production was significantly inhibited when 16HBE cells were stimulated in the presence of sgp130 (Fig. 7 D).

In addition to CCL-2 induction, sIL-6Rα in conjunction with IL-6 decreases CXCL8 production in endothelial, mesothelial, and mesangial cells (7, 9). A significant increase in CXCL8 mRNA levels was observed as early as 1 h after stimulation with S. aureus or P. aeruginosa and 4 h after stimulation with flagella (Fig. 8, A and B), and a significant increase in CXCL8 production was observed 4 h after stimulation (Fig. 7, C and D). By 24 h after S. aureus or flagella stimulation, when high levels of shed IL-6Rα were detected (Fig. 4, A and C), CXCL8 mRNA expression was significantly decreased to the levels found in resting unstimulated cells (Fig. 8, A and B), which correlates with the observation that CXCL8 levels did not increase between 4 and 24 h (Fig. 8, C and D). To demonstrate that shed IL-6Rα was responsible for down-regulating CXCL8 expression, airway epithelial cells expressing pRS-TACE-1 or pRS-sc were stimulated with flagella for different times. Maximum CXCL8 production was observed 4 h after stimulation in cells expressing the vector control, and there was no increase after 24 h of stimulation. Cells lacking TACE expression showed significantly increased levels of CXCL8 after 24-h stimulation compared with that after 4 h, which indicates that shed IL-6Rα is required for CXCL8 down-regulation by airway epithelial cells.

FIGURE 8.

CXCL8 production in epithelial cells. 16HBE cells were stimulated with S. aureus, P. aeruginosa (PAO1), or flagella. A and B, CXCL8 mRNA expression was determined by real-time PCR. Values were normalized to β-actin and are shown as the fold change in expression relative to the endogenous level in unstimulated (C, control) cells. Each bar represents the mean of triplicate samples. ∗, p < 0.01. C and D, CXCL8 production was determined by ELISA. Each bar represents the mean of sextuplicate wells. ∗∗, p < 0.001. E, 16HBE cells expressing pRS-TACE-1 or pRS-sc were stimulated with flagella, and CXCL8 production was determined by ELISA. ∗, p < 0.05.

FIGURE 8.

CXCL8 production in epithelial cells. 16HBE cells were stimulated with S. aureus, P. aeruginosa (PAO1), or flagella. A and B, CXCL8 mRNA expression was determined by real-time PCR. Values were normalized to β-actin and are shown as the fold change in expression relative to the endogenous level in unstimulated (C, control) cells. Each bar represents the mean of triplicate samples. ∗, p < 0.01. C and D, CXCL8 production was determined by ELISA. Each bar represents the mean of sextuplicate wells. ∗∗, p < 0.001. E, 16HBE cells expressing pRS-TACE-1 or pRS-sc were stimulated with flagella, and CXCL8 production was determined by ELISA. ∗, p < 0.05.

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Airway epithelial cells synthesize numerous cytokines, chemokines, antimicrobial peptides, and mucins immediately in response to bacterial ligands and recruit PMNs to the site of infection. Excessive proinflammatory signaling acutely results in airway obstruction and respiratory failure, as frequently occurs in severe bacterial pneumonia. Persistently activated mucosal signaling, a characteristic of cystic fibrosis airway disease, ultimately results in destruction of lung parenchyma and loss of pulmonary function. Thus, the appropriate balance between sufficient pro-inflammatory recruitment to clear an infection and the potentially negative effects of PMN accumulation, release of neutrophil elastase, and toxic oxygen intermediates is particularly critical in the lung.

In the experiments described in this report, we demonstrate that the production of IL-6 and IL-6Rα shedding by epithelial cells have a major role in CCL-2 induction and the switch toward a monocyte/macrophage-dominated response during S. aureus or P. aeruginosa airway infection. IL-6 has been shown to regulate many components of the immune response: modulating gene expression in macrophages (25), affecting C5aR levels (26) and T cell apoptosis (27), as well as the mobilization and activation of PMNs (28). IL-6 signaling through the sIL-6Rα is critical for IFN-γ-mediated PMN infiltration and activation as well as apoptosis (28). The importance of IL-6 in regulating inflammation at mucosal surfaces has been examined in some detail in models of inflammatory bowel disease (29, 30). IL-6 expression by gastrointestinal epithelial cells, induced by a number of physiological stimuli, is sufficient to induce PMN Ca2+ fluxes and stimulate PMN degranulation (31). IL-6/sIL-6Rα trans-signaling contributes to T cell activation in the gut by inhibiting apoptosis and appears to be an important component of mucosal inflammation in Crohn’s disease and experimental colitis (32). The protective role of IL-6 in modulating inflammation has been well documented in models of sepsis (26, 33). The multiple effects of IL-6 on a number of cytokine signaling pathways (IL-1, TNF-α, IL-10, and TGF-β) that are activated by bacterial infection has also been recently documented in a Yersinia enterocolitica model of infection of macrophages (25).

Bacteria not only activate epithelial IL-6 expression immediately upon exposure, but also stimulate transcription of TACE. TACE or ADAM 17 is a member of the ADAM family of proteases, a relatively promiscuous enzyme involved in the release of a number of superficial cellular proteins, including the IL-6, epidermal growth factor (34), and TNFRs (13). TACE is produced as a zymogen and requires processing to remove an inhibitory domain (35). There are little published data regarding the physiological inducers of TACE expression, and most studies use PMA to induce TACE activity (16, 36, 37, 38). There is a c/EBP/NF-IL-6 site upstream of the mouse TACE promoter (39), which is responsive to bacterial stimulation in epithelial cells. Bacterial induction of TACE expression appears to be part of a hierarchy of epithelial responses. CXCL8, IL-6, and TACE transcription are induced upon bacterial contact, and the consequences of TACE activation, the release of sIL-6 Rα and the shedding of surface IL-6Rα, are maximal at 24 h. Epithelial exposure to IL-6 is not sufficient to activate TACE expression, even after prolonged incubation. Bacterial exposure for at least 4 h was required. These results suggest that bacterial ligands are an important physiological activator of TACE expression and activity, which is consistent with its role in controlling the availability of cytokine receptors on the epithelial surface.

TACE activity has additional anti-inflammatory effects on airway cells by regulating responsiveness to TNF-α. TNFR1 is among the substrates for TACE that is present on the apical surface of airway cells. In response to bacterial infection, there is abundant TNF-α present in the airways, produced primarily by immune cells. TACE-dependent cleavage of TNFR1 has been well documented in immune cells (13) as well as on airway epithelial cells in response to S. aureus (40). This TACE-mediated release of TNFR1 serves to neutralize free TNF-α in the airway lumen as well as prevent additional epithelial activation through loss of TNFR1 from the cell.

The consequences of receptor shedding and IL-6/sIL-6Rα trans-signaling that have been detailed in macrophages have not been previously implicated in the control of airway inflammation. Induction of TACE expression in airway cells, followed by loss of surface-associated IL-6Rα, appearance of sIL-6Rα in the cell culture supernatant, induction of CCL-2 expression, and decreased CXCL8 expression appear to be a general response to bacterial stimulation. Airway cells apparently participate in this process, and a regulatory response is initiated at the same time as the initial proinflammatory signaling in response to bacteria. The high affinity binding of the sIL6Rα/IL-6 complex and membrane-associated gp130 occurs on the surface of airway cells. Distal signaling, which involves the formation of a hexameric structure to induce gp130 phosphorylation (41), results in CCL-2 induction and decreased CXCL8 transcription. All the components of this signaling cascade are expressed by airway epithelial cells as a general response to bacterial stimulation.

There are undoubtedly direct affects of leukocyte products on airway cell proinflammatory signaling as well as those initiated by the epithelial cells themselves. Both IL-10 and TGF-β can regulate proinflammatory signals in the airways (4, 5, 6). There is ample evidence of the participation of IL-10 in the control of mucosal inflammation, especially in gastrointestinal mucosa (42). Whether respiratory epithelial cells express IL-10 to self-modulate is not well documented. TGF-β, although clearly important in the resolution of acute airway inflammation, is largely the product of the infiltrating immune cells. What is novel about the involvement of IL-6 signaling in airway cells is that the same initial stimulus, the bacterial ligand, initiates the PMN-dominated proinflammatory signaling and later also stimulates the signaling involved in the switch to a monocyte/macrophage-dominated inflammation.

These studies suggest that mucosal epithelial cells have a primary role in regulating their own signaling capabilities, even in the absence of direction from leukocytes and their products. They are tremendously efficient in responding immediately to bacterial infection by inducing the influx of PMNs and activating them through the induction of CXCL8 and IL-6 expression. By controlling the availability of their own surface receptors through the activity of TACE, airway cells have a mechanism to regulate responses to these endogenously expressed cytokines as well as those produced by the recruited immune cells. Targeting the airway epithelium, as opposed to the systemic immune system, may provide a means to control excessive acute inflammation without the risk associated with systemic immunosuppression.

Confocal microscopy was performed at the Herbert Irving Optical Microscopy facility at Columbia University.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by National Institutes of Health Grants RO1DK39693 and HL56194 and the U.S. Cystic Fibrosis Foundation. M.I.G. was the recipient of a postdoctoral fellowship from the U.S. Cystic Fibrosis Foundation.

4

Abbreviations used in this paper: PMN, polymorphonuclear cell; ADAM, a disintegrin and metalloprotease; ARTS-1, aminopeptidase regulator of TNFR1 shedding; DS, differential mRNA splicing; pRS, pRetroSuper vector; RNAi, RNA interference; s, soluble; sgp130, soluble gp130; TACE, TNF-α converting enzyme;TAPI-1, TNF-α protease inhibitor-1.

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