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Am J Physiol Lung Cell Mol Physiol 290: L86-L96, 2006. First published August 12, 2005; doi:10.1152/ajplung.00391.2004
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Inflammatory response of tracheobronchial epithelial cells to endotoxin

Simona B. Neff,1,3,* Birgit Roth Z'graggen,2,* Thomas A. Neff,1,3 Marina Jamnicki-Abegg,1,2 Dominik Suter,1,2 Ralph C. Schimmer,4 Christa Booy,2 Hana Joch,5 Thomas Pasch,1 Peter A. Ward,3 and Beatrice Beck-Schimmer1,2

1Institute of Anesthesiology and 2Institute of Physiology, University of Zurich Medical School, Zurich, Switzerland; 3Department of Pathology, University of Michigan Medical School, Ann Arbor, Michigan; 4Department of Surgery and 5Cardiovascular Research, Institute of Physiology, University of Zurich Medical School, Zurich, Switzerland

Submitted 21 October 2004 ; accepted in final form 8 August 2005


   ABSTRACT
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 ABSTRACT
METHODS
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 DISCUSSION
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 REFERENCES
 
Respiratory epithelial cells play a crucial role in the inflammatory response in endotoxin-induced lung injury, an experimental model for acute lung injury. To determine the role of epithelial cells in the upper respiratory compartment in the inflammatory response to endotoxin, we exposed tracheobronchial epithelial cells (TBEC) to lipopolysaccharide (LPS). Expression of inflammatory mediators was analyzed, and the biological implications were assessed using chemotaxis and adherence assays. Epithelial cell necrosis and apoptosis were determined to identify LPS-induced cell damage. Treatment of TBEC with LPS induced enhanced protein expression of cytokines and chemokines (increases of 235–654%, P < 0.05), with increased chemotactic activity regarding neutrophil recruitment. Expression of the intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1) was enhanced by 52–101% (P < 0.0001). This upregulation led to increased adhesion of neutrophils, with >95% adherence to TBEC after LPS stimulation, which could be blocked by either ICAM-1 (69%) or VCAM-1 antibodies (55%) (P < 0.05). Enhanced neutrophil-induced necrosis of TBEC was observed when TBEC were exposed to LPS. Reduced neutrophil adherence by ICAM-1 or VCAM-1 antibodies resulted in significantly lower TBEC death (52 and 34%, respectively, P < 0.05). Therefore, tight adherence of neutrophils to TBEC appears to promote epithelial cell killing. In addition to indirect effector cell-induced TBEC death, direct LPS-induced cell damage was seen with increased apoptosis rate in LPS-stimulated TBEC (36% increase of caspase-3, P < 0.01). These data provide evidence that LPS induces TBEC killing in a necrosis- and apoptosis-dependent manner.

lipopolysaccharide; inflammatory mediators; leukocytes

ACUTE LUNG INJURY (ALI) and its progression to the acute respiratory distress syndrome (ARDS) remain leading factors of morbidity and mortality in critically ill patients (14, 20). ARDS is characterized by hypoxemia, high-permeability pulmonary edema, and neutrophil accumulation in the lung. The morphological manifestations may include damage to endothelial and epithelial sites of the blood-gas barrier.

In vitro and in vivo animal studies of lipopolysaccharide (LPS)-induced lung injury are widely used as experimental approaches to investigate the mechanisms of acute lung inflammatory injury. Substantial advances in the understanding of the underlying mechanisms of the inflammatory processes have been made. After LPS administration, the coordinated expression of cytokines [tumor necrosis factor-{alpha} (TNF-{alpha})], CC chemokines [monocyte chemoattractant protein-1 (MCP-1), macrophage inflammatory protein-1{beta} (MIP-1{beta})], CXC chemokines [cytokine-induced neutrophil chemoattractant-1 (CINC-1), macrophage inflammatory protein-2 (MIP-2)], and adhesion molecules [intercellular adhesion molecule-1 (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1)] forms a complex network, which in turn promotes neutrophil extravasation, tissue migration, and, finally, neutrophilic inflammation and tissue damage as part of the injury repair process (54). It is well known that these inflammatory mediators are produced not only by phagocytic cells but also by a variety of structural pulmonary cells (54).

The airway compartment lined with epithelial cells is the predominant structural barrier and the primary interface with pathogens and a variety of environmental agents. Therefore, it represents a pivotal site for the innate immunity and pulmonary host defense. Many studies of the distal airway epithelium, which consists mainly of type I and II alveolar epithelial cells (AEC), have demonstrated its importance in the pathogenesis of and recovery from ALI. It is well known that AEC play a major role in the regulation of the immune and inflammatory responses in the lung (26, 40, 42). However, little information is available about tracheobronchial epithelial cells (TBEC) and their inflammatory responses to LPS (36).

Depending on the cell type and stimulus, cell death may be caused by apoptosis or necrosis. The process of apoptosis is both biochemically and morphologically distinct from necrosis and contributes to primary organ damage (29). Recent reports have indicated that LPS induces disseminated endothelial and alveolar epithelial apoptosis, suggesting that TBEC also might undergo apoptosis upon LPS stimulation (15, 16).

To evaluate the potential of endotoxin to damage epithelial cells of the upper respiratory compartment in vitro, we incubated rat TBEC with LPS. Changes in expression pattern of inflammatory mediators were determined as well as the biological consequences regarding epithelial cell necrosis and apoptosis. The present study was initiated to determine whether, in addition to AEC, TBEC also are involved in the inflammatory response to LPS and to assess the respective mechanisms.


   METHODS
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Animals. Specific pathogen-free male Wistar rats (250–300 g) were purchased from Janvier (Le Genest-St. Isle, France). Rats were anesthetized with subcutaneously administered Hypnonorm (fentanyl/fluonisone; Janssen, Beerse, Belgium) at 0.25 ml/kg body wt, Domitor (medetomidine; Pfizer, Westchester, PA) at 0.25 ml/kg body wt, and atropine at 0.05 ml/kg body wt. All animal experiments and animal care were approved by the Swiss Veterinary Health Authorities.

Isolation and culture of TBEC. Trachea and bronchial parts of the respiratory system were excised, ligated at the distal ends, filled with 0.01% protease type XIV (Sigma, Buchs, Switzerland), and incubated overnight at 4°C. TBEC were retrieved using fetal bovine serum (FBS) and washed twice. They were then incubated in airway epithelial cell basal medium with supplements (PromoCell, Heidelberg, Germany)-10% premium FBS (BioWhittaker, Verviers, Belgium)-1% penicillin-streptomycin in 35-mm culture plates precoated with 50 µg/ml rat tail collagen (Sigma) for 30 min at room temperature. The medium was supplemented with 0.4% bovine pituitary extract, 0.5 ng/ml epidermal growth factor, 5 µg/ml insulin, 0.5 µg/ml hydrocortisone, 0.5 µg/ml epinephrine, 6.7 ng/ml triiodothyronine, 10 µg/ml transferrin, and 0.1 ng/ml retinoic acid. Under these conditions, cells grew and reached 100% confluence within 3 days. Purity was verified using periodic acid-Schiff staining (>98%). Epithelial cell character also was confirmed by a pathologist at the University Hospital of Zurich.

Isolation and culture of AEC. AEC were harvested according to an established protocol (3). Briefly, lungs were incubated with porcine pancreas elastase. Enzyme activity was stopped with FBS, and tissue was then minced. Filtered cells were incubated in IgG-coated plastic plates, and unattached cells were collected after 1 h. Cells were added to culture plates, and experiments were performed when cells were confluent. Purity was assessed with a fluorescein-labeled lectin, Griffonia simplicifolia I (macrophage staining), because macrophages were the critical contaminating cell population. Cell purity was always >95%.

Stimulation with LPS. Confluent monolayers of TBEC were stimulated with LPS (Escherichia coli, serotype 055:B5; Sigma) at 100 µg/ml as previously described for AEC (3, 26). Dose-response experiments showed that the same inflammatory response was seen between 5 and 100 µg/ml LPS. However, to be consistent with earlier experiments (3, 26), we used LPS at a concentration of 100 µg/ml.

RT-PCR for TNF-{alpha}, MCP-1, MIP-1{beta}, CINC-1, MIP-2, ICAM-1, and VCAM-1-mRNA. Total RNA was extracted from confluent monolayers of unstimulated (control) and LPS-stimulated TBEC by using Trizol (Life Technologies, Basel, Switzerland) according to the manufacturer's instructions. Total RNA (0.8 µg) was reverse transcribed, and PCR was performed with primers for rat TNF-{alpha}, MCP-1, MIP-1{beta}, CINC-1, MIP-2, ICAM-1, and VCAM-1 as described in Table 1. PCR also was performed with 18S primers. The PCR product was confirmed by electrophoresis in a 1.5% agarose gel. For quantitative analysis of PCR, densitometry was performed for mRNA of the inflammatory mediators and 18S. mRNA/18S ratios were calculated. One of the five control values (PBS) was defined as 100, and all others were normalized.


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  		Table 1. Optimized conditions for RT-PCR

 
Enzyme-linked immunosorbent assay quantification of TNF-{alpha}, MCP-1, CINC-1, and MIP-2 in supernatant fluids of LPS-stimulated TBEC. TNF-{alpha} and MCP-1 were assessed in the supernatant fluids of LPS-stimulated TBEC by using standard enzyme-linked immunosorbent assay (ELISA) purchased from BD Biosciences (San Diego, CA); rat CINC-1 and MIP-2 were assessed using an ELISA purchased from R&D Systems (Abingdon, UK). The minimum detectable concentration of proteins was 15 pg/ml for TNF-{alpha}, 31.3 pg/ml for MCP-1, 15.6 pg/ml for CINC-1, and 31.2 pg/ml for MIP-2.

Western blot analysis of MIP-1{beta} in supernatant fluids of LPS-stimulated TBEC. Supernatant fluids of LPS-stimulated TBEC were loaded and electrophoresed in a 12.5% SDS-polyacrylamide gel. After separation, the proteins were transblotted to a nitrocellulose membrane for 2 h at 200 mA (Bio-Rad, Hercules, CA). Equal loading of proteins was confirmed by Ponceau S staining. The blot was washed in PBS and blocked with PBS-4% low fat milk-0.1% Tween 20 for 1 h at room temperature, followed by an overnight incubation with a polyclonal rabbit anti-rat MIP-1{beta} antibody (Biovision, Lausen, Switzerland) diluted 1:100 in blocking buffer. All washing steps were performed three times with PBS-0.1% Tween 20. A secondary horseradish peroxidase-labeled anti-rabbit IgG (Sigma) diluted 1:5,000 in blocking buffer was added for 1 h at room temperature. Signals were detected by enhanced chemiluminescence.

ELISA for ICAM-1 and VCAM-1 expression. To determine ICAM-1 and VCAM-1 expression on the surface of TBEC, we performed a cell-based ELISA according to a previous protocol (4, 26). ICAM-1 antibody was a monoclonal mouse anti-rat antibody (Serotec, Oxford, UK), and VCAM-1 antibody was a polyclonal goat anti-rat antibody (Santa Cruz Biotechnology, Santa Cruz, CA). Final optical density (OD) was calculated by subtracting the OD of cells incubated with the secondary antibody alone or with a control antibody (MOPC-21, mouse IgG; goat IgG) (Pharmingen, San Diego, CA) from the OD of cells incubated with primary and secondary antibody together.

In vivo model of LPS-induced lung injury. Anesthetized animals were exposed to intratracheally instilled LPS (150 µg in 300 µl of PBS; 300 µl of PBS alone for control animals) (3). Lungs were subsequently analyzed at predefined time points. Lungs were lavaged with 10 ml of cold PBS, with instillation and aspiration repeated five times.

Tissue sampling by laser capture microdissection: preparation of RNA and cDNA for VCAM-1. Lungs were inflated with 10 ml of Tissue-Tek (MICROM International, Walldorf, Germany) and snap frozen. Frozen sections of 6-µm thickness were obtained and mounted on a membrane (Molecular Machines Industries, Glattbrugg, Switzerland). Sections were semi-dried for 10 s, followed by fixation in 70% ethanol for 30 s, twice in 95% ethanol for 15 s, twice in 100% ethanol for 15 s, and twice in xylene for 1 min. Finally, sections were dried for 4 min. Before laser capture microdissection (LCM), the membranes were thawed and completely dried to prevent RNase activity. LCM was performed on an inverse microscope (Nikon TE300 with laser microdissection) at a x400 magnification. A total of 40–60 pieces of bronchial epithelial tissue with a total area of ~500,000 µm2 were collected on each cap of a microtube (Molecular Machines Industries). The captured cells were dissolved in lysis buffer. Total RNA was extracted from the laser-captured cells by using the Absolutely RNA Nanoprep kit according to the manufacturer's directions (Stratagene, La Jolla, CA). RT-PCR was performed using a TaqMan RT-PCR kit (TaqMan reverse transcription reagents; PE Applied Biosystems, Foster City, CA). A total volume of 30 µl, containing 37.5 units of MultiScribe reverse transcriptase (PE Applied Biosystems) and 2.5 µM random hexamers (PE Applied Biosystems), was incubated at 25°C for 10 min, at 48°C for 30 min, and at 95°C for 5 min. The resulting cDNA was used for the quantitative PCR (QPCR) as described below.

Real-time TaqMan PCR analysis. VCAM-1 gene expression was analyzed using a quantitative real-time PCR procedure with an ABI Prism 7700 sequence detection system (PE Applied Biosystems). PCR primers were designed using Primer Express software (PE Applied Biosystems).

QPCR was performed using Brilliant SYBR green PCR mastermix (Stratagene Europe; no. 600548) according to the manufacturer's protocol. Briefly, the reaction contained 12.5 µl of SYBR green mastermix, 2.5 µl of primer sense, 2.5 µl of primer-antisense (150 nM VCAM, 50 nM 18S), 0.37 ml of the reference dye Rox diluted at 1:50 (Stratagene), and 3 µl of cDNA. After the activation step for Taq polymerase at 95°C for 10 min, PCR was started using the following cycling parameters: denaturation at 95°C for 30 s, annealing at 60°C for 1 min, and extension at 72°C for 1 min, 40 cycles.

QPCR reactions for VCAM-1 and 18S were performed in separate tubes to avoid possible competition and/or interference in a single reaction tube. The following oligonucleotide primers (5'-3' sequences) were used for QPCR analysis: VCAM-1 sense, 5'-AAAGAAAGGAGACTGTCAGAGAACT-3'; VCAM-1 antisense, 5'-CTTGTGGAGGGATGTACAGAGATCT-3'; 18S sense, 5'-TGAGGCCATGATTAAGAGGG-3'; and 18S antisense, 5'-AGTCGGCATCGTTTATGGTC-3'.

Gene expression was normalized to the 18S, and the increase in VCAM-1 gene expression was calculated with the comparative CT (threshold cycle) method of gene expression in stimulated and unstimulated bronchial epithelial cells. VCAM-1 gene expression in unstimulated samples was assigned a baseline value of 1.

Preparation of alveolar macrophages and neutrophils. Rat alveolar macrophages were harvested according to a well-established protocol (52). Human neutrophils were obtained from healthy human volunteers and isolated as described previously (12, 26). The use of human neutrophils for rat assays is well established (41), and experiments with rat neutrophils have been proven to produce identical results compared with the use of human neutrophils in our laboratory. Because it was impossible to retrieve a sufficient number of rat neutrophils for all the proposed experiments, human neutrophils were preferred. However, it always has to be kept in mind that neutrophils in this experimental setup came from a different species.

Chemotaxis assay. Neutrophils [5 x 106 polymorphonuclear neutrophils (PMN)/ml] were labeled at a concentration of 5 µM with the fluorescent indicator calcein acetoxymethyl ester (calcein-AM; Calbiochem Biochemicals, Juro Supply, Lucerne, Switzerland) for 30 min at 37°C. PMN were subsequently washed twice with PBS. Labeled neutrophils (1 x 105 PMN) were placed into MultiScreen-MIC filter plates (Millipore, France). Receiver plates were loaded with 150 µl of supernatant of previously treated cells as described earlier (24-h LPS stimulation). The chemotactic substance N-formyl-methionyl-leucyl-phenylalanine (0.1 µM; Sigma) was taken as positive control, and DMEM-1% FBS was used to determine basal migration. After 2-h incubation at 37°C, the filter plate was taken out and migrated cells were lysed with 1.0% Triton-X (Sigma). Fluorescence was measured using an excitation filter at 485 nm and an emission filter at 535 nm. The ratio between migration to supernatant of stimulated TBEC and basal migration was used as an indication of PMN chemotaxis.

Assay of alveolar macrophage and neutrophil adherence to TBEC. TBEC were added to 96-well plates at a density of 5 x 104 cells/well and grown to confluence at 37°C in 5% CO2. Cells were stimulated with 100 µg/ml LPS for 24 h. After the cells were washed, 1 x 105 alveolar macrophages or neutrophils were added to each well and incubated at 37°C in 5% CO2 for 30 min. Fifteen minutes later, phorbol 12-myristate 13-acetate (PMA; Sigma) was added to some wells to stimulate effector cells (25 ng/ml final concentration). Nonadherent cells were removed by washing each well four times. Remaining macrophages or neutrophils were counted immediately.

For blocking experiments, neutrophils were preincubated at 37°C for 15 min with both the monoclonal mouse anti-human LFA-1/CD11a and Mac-1/CD11b antibodies (10 µg/ml) or with monoclonal mouse anti-human CD49d antibody (10 µg/ml) (Becton Dickinson, Basel, Switzerland) (26). For controls, a nonspecific binding monoclonal mouse anti-human CD40 antibody (10 µg/ml) (Becton Dickinson) was used. Before their addition to the TBEC monolayers, neutrophils were labeled with calcein AM. Neutrophils were then added to each well for 30 min and incubated at 37°C in 5% CO2 as described above. Nonadherent PMN were removed by carefully washing the cells twice with DMEM. DMEM (100 µl) was added to each well before fluorescence measurement, and fluorescence was measured using an excitation filter at 485 nm and an emission filter at 535 nm. The amount of adherent PMN was calculated using a standard row.

Cytotoxicity assay. Effector cell-induced cytotoxicity was determined by measuring release of 51Cr from epithelial cells into the supernatant fluids (49, 50). Epithelial cells were cultured in 96-well plates. 51Cr (1 µCi) was added to each well and incubated for 24 h. To stimulate epithelial cells, we added LPS for 24 h (PBS for control cells). Macrophages or neutrophils, previously diluted in PBS-0.02% bovine serum albumin, were added to the epithelial cells and stimulated with PMA as described. Effector cells were incubated with target cells for 2, 4, and 6 h. Supernatant fluids were centrifuged, and 51Cr remaining in the supernatant fluids was measured. Some TBEC were incubated with a lysing solution (1% Triton-X) for 45 min (total 51Cr release). Supernatant fluids also were collected from wells of TBEC, which were not incubated with alveolar macrophages/neutrophils (spontaneous 51Cr release). Cytotoxicity was calculated according to the following formula: cytotoxicity (%) = (ER – SR)/(TR – SR) x 100, where ER is experimental release, SR is spontaneous release, and TR is total release. The same experiments were performed with AEC as target cells, using alveolar macrophages and neutrophils.

For blocking experiments with ICAM-1 and VCAM-1 antibodies, effector cells were preincubated with LFA-1/CD11a and Mac-1/CD11b antibodies, with CD49d antibody, or with CD40 antibody as described above. The cytotoxicity assay was performed according to the described protocol.

Fluorometric assay for caspase-3 and caspase-8 activity. For the caspase assays, TBEC were isolated and grown to confluence in 96-well plates and stimulated with LPS (100 µg/ml) or camptothecin (4 µM) for 24 h. Caspase-3 and caspase-8 activity was determined by measuring proteolytic cleavage of the fluorogenic caspase-3 substrate Ac-Asp-Glu-Val-Asp-AMC (Merck, Darmstadt, Germany) or caspase-8 substrate Z-Ile-Glu-Thr-Asp-AFC (Calbiochem). Cells were incubated for 1 h at 37°C with 125 µM substrate. The fluorescence of the cleaved reporter group was measured at an excitation wavelength of 360 nm and an emission wavelength of 465 nm.

Camptothecin (an extract of the Chinese tree Camptotheca acuminata) is a potent inhibitor of topoisomerase I, a molecule required for DNA synthesis. Camptothecin has been shown to induce apoptosis and was therefore used for positive controls (47).

Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling assay. TBEC were grown in eight-well glass chamber slides (VWR International, Dietikon, Switzerland). Cells were stimulated with LPS and camptothecin, fixed in 4% paraformaldehyde, and washed with PBS-0.1% Triton. Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) was carried out according to the instructions provided by the manufacturer (Terminal Transferase, recombinant; Roche, Basel, Switzerland; fluorescein-12–2'-deoxyuridine-5'-triphosphate, Enzo; Roche Diagnostics, Penzberg, Germany).

Preparation of cytosolic protein fraction. Confluent TBEC stimulated for 24 h with or without LPS were harvested and washed three times in Ca2+- and Mg2+-free Hanks' balanced salt solution. Two milliliters of homogenization buffer (0.5 µg/ml protease inhibitor mix, 1 mM PMSF, 250 mM sucrose, 1 mM EGTA, 1 mM EDTA, and 10 mM HEPES/NaOH, pH 7.4) were added, and the pellets were homogenized on ice. Homogenate was centrifuged (1,500 g, 10 min, 4°C), and supernatant was transferred into a clean tube. This step was repeated twice, and supernatants were combined and centrifuged again (1,500 g, 10 min, 4°C). With the obtained supernatant centrifugation was performed at much higher speed (10,000 g, 12 min, 4°C). The pellet was discharged, and supernatant was centrifuged for 10 min at 16,000 g, with the resulting supernatant containing the cytosolic fraction of proteins. Protein concentrations were determined using the Bradford protein assay.

Determination of cytosolic cytochrome c. Cytosolic proteins (10 µg/lane) were separated electrophoretically on a 15% SDS-polyacrylamide gel and transferred to a nitrocellulose membrane as described. Purified 15 kDa cytochrome c (Sigma) was used as control (3 ng). The membrane was stained with Ponceau S to confirm uniform protein loading and to ensure equal amounts of protein being transferred to the membrane. Monoclonal mouse antibody directed against cytochrome c (1:500; BD Pharmingen, Basel, Switzerland) was incubated overnight, followed by an incubation with a secondary antibody (goat anti-mouse IgG conjugated to horseradish peroxidase, 1:10,000).

Rat cytochrome c immunoassay. Cytochrome c ELISA kit (R&D Systems, Oxon, UK) was used to assess the protein content of cytochrome c in the mitochondria-free cytosolic fraction of TBEC to evaluate the release of the mitochondrial cytochrome c to the cytosol. Following the manufacturer's protocol, we used 50 µl of the extracted cytosolic fraction as an antigen source in a sandwich ELISA. The change in color was monitored at a wavelength of 450 nm with the use of a Multiscan RG plate reader. Measurements were performed with the control and suspended samples analyzed on the same microplate, and cytochrome c content was expressed as nanograms per milliliter of cytochrome c compared with the standard curve.

Statistical analysis. All experiments were repeated at least three times. Results are expressed as means ± SE and t-test (unpaired, two-tailed). Statistical significance of the differences between means was determined at a 5% level. Densitometry was performed with n = 5. Statistical calculations were performed using GraphPad Prism 2.0 computer software (San Diego, CA).


   RESULTS
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 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Chemokine and cytokine production of TBEC after stimulation with LPS. The potential release of monocyte or neutrophil chemoattractants into the supernatant upon LPS stimulation was investigated for several inflammatory mediators. Figure 1A shows mRNA levels for TNF-{alpha}, MCP-1, MIP-1{beta}, CINC-1, and MIP-2 in unstimulated and LPS-stimulated TBEC. LPS stimulation led to an increase of TNF-{alpha} mRNA by 67% (P < 0.05), MCP-1 mRNA by 37% (P < 0.05), MIP-1{beta} mRNA by 27% (P < 0.01), CINC-1 mRNA by 96% (P < 0.05), and MIP-2 mRNA by 53% (P < 0.05). After LPS stimulation, TNF-{alpha} protein concentration in the supernatant fluids increased by 335% (P < 0.01), MCP-1 by 326% (P < 0.05), MIP-1{beta} by 654% (P < 0.0001), CINC-1 by 235% (P < 0.057, not significant), and MIP-2 by 349% (P < 0.05) (Fig. 1B).



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  		Fig. 1. A: tumor necrosis factor-{alpha} (TNF-{alpha}), monocyte chemoattractant protein-1 (MCP-1), macrophage inflammatory protein-1{beta} (MIP-1{beta}), cytokine-induced neutrophil chemoattractant-1 (CINC-1), and macrophage inflammatory protein-2 (MIP-2) mRNA levels in tracheobronchial epithelial cells (TBEC): unstimulated cells (control, open bars) and cells stimulated (solid bars) with Escherichia coli (serotype 055:B5) lipopolysaccharide (LPS; 100 µg/ml) for 24 h. Total cellular mRNA was extracted, and TNF-{alpha}, MCP-1, MIP-1{beta} CINC-1, and MIP-2 mRNA was detected by RT-PCR. Equal loading was shown with 18S bands. Densitometry was performed in relation to 18S (n = 5). Data show means from a representative experiment. *P < 0.05; **P < 0.01 compared with unstimulated TBEC. B: levels of TNF-{alpha}, MCP-1, MIP-1{beta}, CINC-1, and MIP-2 released into supernatant fluids of TBEC. TNF-{alpha}, MCP-1, CINC-1, and MIP-2 protein determination was performed with ELISA, whereas MIP-1{beta} protein determination was performed with Western blotting (left lane, unstimulated cells; right lane, LPS-stimulated TBEC). Densitometry was performed with 5 different experiments (only 1 blot is shown). TNF-{alpha}, MCP-1, CINC-1, and MIP-2 ELISAs and MIP-1{beta} Western blot: unstimulated cells (control) and cells after stimulation with LPS (100 µg/ml) for 24 h. Data show means ± SE from a representative experiment with n = 5. *P < 0.05; **P < 0.01; ***P < 0.0001 compared with unstimulated TBEC.

 
Chemotaxis assay. Neutrophils are main effector cells in acute lung injury. To assess the biological function of the chemokines released from TBEC regarding neutrophil recruitment, we performed chemotaxis assays. The results showed a 370% (P < 0.0001) increase of migrated PMN to the supernatant of LPS-stimulated cells compared with the control group incubated with PBS. (Fig. 2).



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  		Fig. 2. Determination of chemotactic activity in supernatants of TBEC. Calcein-AM-labeled polymorphonuclear neutrophils (PMN; 1 x 105) were placed into MultiScreen-MIC filter plates, and receiver plates were loaded with 150 µl of supernatant of TBEC, pretreated with LPS (100 µg/ml) or PBS. DMEM-1% FBS was used to determine basal migration. After a 2-h incubation fluorescence was measured using an excitation filter at 485 nm and an emission filter at 535 nm. The ratio between migration to supernatant of stimulated L2 cells and basal migration was used as an indication of PMN chemotaxis. Values are means ± SE from 5 experiments. *P < 0.0001

 
Effects of LPS on mRNA and protein expression for ICAM-1 and VCAM-1 in TBEC. The adhesion molecules ICAM-1 and VCAM-1 are important mediators in many inflammatory processes. Therefore, the expression patterns for these adhesion molecules were defined. TBEC were stimulated with LPS (100 µg/ml) for 24 h, and total RNA was extracted. A representative result is shown in Fig. 3A. LPS-stimulated TBEC showed a slight increase in ICAM-1 mRNA expression, compared with unstimulated cells (35% increase, P < 0.05). However, an upregulation of 105% occurred for VCAM-1 mRNA after stimulation with LPS (P < 0.0001). To determine ICAM-1 and VCAM-1 protein expression in TBEC, we used a cell-based ELISA. LPS induced an increase in ICAM-1 expression by 52% (P < 0.0001) and in VCAM-1 by 101% (P < 0.0001) (Fig. 3B).



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  		Fig. 3. A: intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1) mRNA levels in TBEC: unstimulated cells (control, open bars) and cells after stimulation (solid bars) with LPS (100 µg/ml) for 24 h. Total cellular mRNA was extracted, and ICAM-1 and VCAM-1 mRNA was detected by RT-PCR. Equal loading was shown with 18S bands. Densitometry was performed in relation to 18S (n = 5). Data show means from a representative experiment. *P < 0.05; **P < 0.0001 compared with unstimulated TBEC. B: cell-based ELISA for ICAM-1 and VCAM-1 protein expression in TBEC: unstimulated cells (control) and cells after stimulation with LPS (100 µg/ml) for 24 h. Data show means ± SE from a representative experiment with n = 5. **P < 0.0001 compared with unstimulated TBEC.

 
In vivo upregulation of VCAM-1 in TBEC after endotracheal stimulation with LPS. Because previous studies suggested that ICAM-1 is upregulated in TBEC in vivo upon LPS stimulation, we were interested to evaluate whether bronchial epithelial VCAM-1 mRNA is enhanced as well in LPS-injured rat lungs (5). An enhancement of VCAM-1 mRNA of 279% was confirmed 24 h after airway instillation of LPS in bronchial cells (Fig. 4). Therefore, in vitro results are comparable with these in vivo data.



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  		Fig. 4. Quantitative TaqMan PCR analysis for VCAM-1 transcript expression of laser-captured bronchial epithelial cells in control lungs and in lungs stimulated with LPS (150 µg intratracheally applied) for 24 h. Data show values and mean from 6 animals.

 
Adherence of alveolar macrophages and neutrophils to LPS-stimulated TBEC. In addition to the demonstration of enhanced adhesion molecule expression, their biological function was explored. Without PMA activation of effector cells, adhesion of alveolar macrophages increased by 554% (P < 0.0001) after TBEC exposure to LPS (Fig. 5A), whereas neutrophil adherence was enhanced by 122% (P < 0.05; Fig. 5B). With PMA stimulation of alveolar macrophages and neutrophils, adherence of alveolar macrophages to LPS-stimulated TBEC increased by 32% (P < 0.05; Fig. 5A). PMA-activated neutrophil adherence to TBEC, however, showed an increase of 95% after stimulation with LPS (P < 0.05; Fig. 5B).



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  		Fig. 5. A: adherence of alveolar macrophages to TBEC. TBEC were unstimulated (control, open bars) or stimulated (solid bars) with LPS (100 µg/ml) for 24 h, and alveolar macrophages [unstimulated or phorbol 12-myristate 13-acetate-(PMA) stimulated] were incubated with TBEC. Nonadherent cells were removed, and remaining macrophages were counted. Data show means ± SE from a representative experiment with n = 5. *P < 0.05; **P < 0.0001 compared with unstimulated TBEC. B: adherence of neutrophils to TBEC. TBEC were unstimulated (control) or stimulated with LPS (100 µg/ml) for 24 h, and neutrophils (unstimulated or PMA stimulated) were incubated with TBEC. Nonadherent cells were removed, and remaining neutrophils were counted. Data show means ± SE from a representative experiment with n = 5. *P < 0.05 compared with unstimulated TBEC. C: ICAM-1- and VCAM-1-dependent adherence of neutrophils to TBEC. TBEC were unstimulated (control) or stimulated with LPS (100 µg/ml) for 24 h. Neutrophils were preincubated with LFA-1/CD11a and Mac-1/CD11b antibody, with CD49d antibody, or with CD40 antibody as control antibody. Calcein-AM-labeled PMN (5 x 105) were added to each well for 30 min. Nonadherent PMN were removed, and fluorescence was measured using an excitation filter at 485 nm and an emission filter at 535 nm. The amount of adherent PMN was calculated using a standard row. Data show means ± SE from a representative experiment with n = 5. *P < 0.05 between LPS group with ICAM-1 or VCAM-1 blocking and LPS group with CD40.

 
ICAM-1- and VCAM-1-mediated neutrophil adhesion was elucidated by indirectly blocking these two cell surface molecules with antibodies to the respective receptor on neutrophils (LFA-1/CD11a and Mac-1/CD11b for ICAM-1, CD49d for VCAM-1) (Fig. 5C). ICAM-1-mediated adhesion could be blocked by 69% (P < 0.05) and VCAM-1 by 55% (P < 0.05).

Damage to TBEC caused by alveolar macrophages and neutrophils. To further investigate the biological impact of increased effector cell adhesion, we examined the cytotoxic potential of alveolar macrophages and neutrophils on TBEC. PMA-stimulated alveolar macrophages were incubated with unstimulated or LPS-stimulated TBEC for 2–6 h. For any time point, no difference in alveolar macrophage-induced cytotoxicity was seen between LPS-stimulated and unstimulated TBEC (not shown). Figure 6A shows a chromium assay to determine cytotoxicity of TBEC incubated for different time points with PMA-stimulated neutrophils. After 2 and 4 h, cytotoxicity was significantly increased in LPS-stimulated TBEC compared with unstimulated controls (2 h: 0.8% in control and 6.2% in stimulated cells, P < 0.05; 4 h: 11.7% in control and 18.8% in stimulated cells, P < 0.01). At later time points (6 h), no difference was detectable between unstimulated and stimulated TBEC.



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  		Fig. 6. A: TBEC killing by PMA-stimulated neutrophils, measured as 51Cr release from TBEC into supernatant fluid. TBEC were unstimulated (control, open bars) or stimulated (solid bars) with LPS (100 µg/ml) for 24 h, followed by incubation with PMA-stimulated neutrophils for 2–6 h. Data show means ± SE from a representative experiment with n = 5. *P < 0.05; **P < 0.01 compared with unstimulated TBEC. B: alveolar epithelial cell (AEC) killing by PMA-stimulated neutrophils, measured as 51Cr release from AEC into supernatant fluid. AEC were unstimulated (control) or stimulated with LPS (100 µg/ml) for 24 h, followed by incubation with PMA-stimulated neutrophils for 2–6 h. Data show means ± SE from a representative experiment with (n = 5). **P < 0.01; ***P < 0.0001 compared with unstimulated TBEC. C: effect of ICAM-1 and VCAM-1 blockade on TBEC killing by PMA-stimulated neutrophils, measured as 51Cr release from TBEC into supernatant fluid. TBEC were unstimulated (control) or stimulated with LPS (100 µg/ml) for 24 h, followed by incubation with PMA-stimulated neutrophils for 4 h, previously treated with LFA-1/CD11a and Mac-1/CD11b antibody, with CD49d antibody, or with CD40 antibody as control antibody. Data show means ± SE from a representative experiment with n = 5. *P < 0.05 between LPS group with VCAM-1 blocking and LPS group with CD40. **P < 0.01 between LPS group with ICAM-1 blocking and LPS group with CD40 or between PBS group with ICAM-1 or VCAM-1 blocking and PBS group with CD40.

 
The same cytotoxicity experiments were performed with AEC as target cells to compare cell necrosis of the upper respiratory compartment with necrosis in the lower one. Again, no AEC killing was seen with alveolar macrophages (not shown). PMA-stimulated neutrophils induced the following increases in cytotoxicity: after 4 h, 20.1% in control and 42.0% in stimulated cells (P < 0.0001); after 6 h, 39.5% in control and 60.5% in stimulated cells (P < 0.01) (Fig. 6B).

We further questioned the importance of firm neutrophil adherence for cytotoxicity. To elucidate this problem, we performed ICAM-1 and VCAM-1 blocking studies. Cytotoxicity was decreased by 52% (P < 0.01) under ICAM-1 blockade, whereas VCAM-1 blockade reduced cytotoxicity by 34% (P < 0.05) (Fig. 6C).

Apoptosis. Cytotoxicity assays revealed evidence for enhanced necrosis rates upon LPS stimulation. However, no information is available about the direct effect of LPS on apoptosis in TBEC. Therefore, caspase-3 activity was determined and TUNEL tests were performed. After TBEC were stimulated with LPS, caspase-3 activity increased by 36% (P < 0.01) (Fig. 7A). TUNEL tests further confirmed an increased apoptosis rate upon LPS stimulation as shown in Fig. 7B: although no apoptotic cells were detectable in control TBEC (a), enhanced nick-end labeling was found in LPS-stimulated TBEC (b). As a positive control, camptothecin was used (c).



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  		Fig. 7. A: caspase-3 assay in TBEC: unstimulated cells (control) and cells after stimulation with LPS (100 µg/ml) for 24 h. Fluorescence of cleaved substrates was determined using spectrophotometry. Data show means ± SE from a representative experiment with n = 5. *P < 0.01 compared with unstimulated TBEC. B: immunofluorescence staining of apoptotic TBEC by terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling method: a, unstimulated cells (control); b, cells after stimulation with LPS (100 µg/ml); c, positive control for apoptosis (TBEC stimulated with 4 µM camptothecin). Apoptotic cells show bright fluorescence. Magnification, x40.

 
To further focus on apoptosis pathway in LPS-injured TBEC, we determined caspase-8 activity, finding no difference between control and LPS-stimulated TBEC (Fig. 8A). However, evaluation of the mitochondrial pathway revealed evidence for an apoptosis pathway with cytochrome c involved. Immunoblotting of TBEC cytosol for cytochrome c showed a single band of ~15 kDa in control and stimulated TBEC (Fig. 8B). At 24 h of LPS stimulation, cytochrome c was significantly augmented in the cytosolic fraction compared with untreated TBEC. These results were verified with the help of an immunoassay, showing an increase of 162% (P < 0.0001) (Fig. 8C).



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  		Fig. 8. A: caspase-8 assay in TBEC: unstimulated cells (control, open bar) and cells after stimulation (solid bar) with LPS (100 µg/ml) for 24 h. Caspase-8 activity was also determined in a positive control (shaded bar), provided by the manufacturer of the kit. Fluorescence of cleaved substrates was determined using spectrophotometry. Data show means ± SE from a representative experiment with n = 5. B: protein immunoblotting for cytochrome c in TBEC: unstimulated cells (lane 2) and cells after stimulation (lane 3) with LPS (100 µg/ml) for 24 h. To ensure equal loading, Ponceau S staining was performed. As a control, lane 1 was loaded with purified 15-kDa cytochrome c. One representative experiment is shown for n = 5. C: rat cytochrome c immunoassay: unstimulated cells (control) and cells after stimulation with LPS (100 µg/ml) for 24 h. Cytosolic, mitochondria-free fraction was extracted, and cytochrome c was detected by ELISA. Data show means ± SE from a representative experiment with n = 5. *P < 0.0001 compared with unstimulated TBEC.

 

   DISCUSSION
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The main goal of this study was to investigate the role of TBEC in the host defense to LPS exposure in vitro. Increased production of proinflammatory cytokines, chemokines, and adhesion molecules in primary culture of rat TBEC in response to in vitro stimulation with LPS was found to significantly increase chemotactic activity for neutrophils. Concomitantly, enhanced ICAM-1- and VCAM-1-dependent adherence of neutrophils induced TBEC killing in cells previously exposed to LPS. In addition to effector cell-induced epithelial cell necrosis, the process of apoptosis also was involved in LPS-induced injury of the cells of the upper respiratory compartment.

LPS are the major outer surface membrane components present in gram-negative bacteria and are strong stimulators of the innate immunity (38). The interaction of the lipid A structure, which imparts the biological activity of LPS, with pattern-recognition receptors results in cellular activation (38). LPS may activate vascular endothelial cells, smooth muscle cells, and fibroblasts, as well as epithelial cells (17, 25, 37). There is increasing evidence that the alveolar epithelium is an important cellular component participating in many stages of ALI (42). In vitro, primary cultures of rat AEC have been shown to secrete inflammatory mediators in response to LPS or endogenous factors such as interleukin-1{beta} (IL-1{beta}) and TNF-{alpha} (9, 31, 34). ICAM-1, which is known to be an important adhesion molecule for neutrophil accumulation, was significantly upregulated on AEC after stimulation with LPS in vitro and in vivo (3). Skerrett et al. (43) recently provided evidence for an active role of the distal airway compartment in the LPS-induced pulmonary inflammation by selectively blocking nuclear factor-{kappa}B (NF-{kappa}B) activation in the distal airway epithelium in vivo. However, less information exists about the upper respiratory epithelial compartment and its involvement in the inflammatory response to LPS.

The fully differentiated normal bronchial epithelium of the larger airways is a stratified structure consisting of a columnar layer comprising ciliated and secretory cells supported by basal cells. This mucociliary epithelium fulfills many functions, which are essential for maintaining the integrity of the respiratory tract. One of the critical functions of the upper respiratory tract is to produce secretions that protect the airways against microbial, particulate, and chemical toxins that contaminate the breathing air. In the smaller airways, bronchial epithelial cells have been speculated to be a key regulator of airway biology in various inflammatory pulmonary diseases, e.g., allergic asthma, chronic obstructive pulmonary disease, and cystic fibrosis (6, 19, 35, 45). These functions are clearly different from AEC, which are important for oxygen transport, production of surfactant, and, upon injury, proliferation of type II into type I alveolar epithelial cells.

The findings of inflammatory response to LPS in TBEC in vitro are controversial: although some authors did not find a response of TBEC to LPS in vitro (10, 28, 33), others have described the release of inflammatory mediators upon LPS stimulation (23, 24). A possible explanation for controversial data in studies performed in these cells might be the change of morphology in TBEC after in vitro culture, with them no longer being ciliated. The study of Koyama et al. (23) showed increased chemotactic activity in supernatant fluids of LPS-stimulated bovine bronchial epithelial cells. The specific chemoattractants, however, have not been directly identified so far. Our findings thus support and expand on these earlier investigations indicating a role for bronchial epithelial cells in regulating the pulmonary inflammatory response to LPS by chemoattractants: not only TNF-{alpha} but also MCP-1, MIP-1{beta}, CINC-1, and MIP-2 are upregulated in TBEC upon LPS stimulation.

MCP-1 is produced in vitro by various epithelial cells, including AEC, in response to inflammatory stimuli such as IL-1{beta}, TNF-{alpha}, and LPS (34). Although its contribution to acute lung injury was initially questioned (7), it was later demonstrated in vivo that the joint presence of MCP-1 and endotoxin in the alveolar compartment can synergize to cause a dramatically enhanced lung inflammatory response or play a major role in neutrophil recruitment (30, 44).

The chemokine MIP-1{beta} plays a significant role in the induction and recovery of acute inflammatory reactions in the lung (7). MIP-1{beta} is a potent chemoattractant for monocytes and lymphocytes (39, 46, 51). Furthermore, it stimulates the production of other inflammatory mediators such as TNF-{alpha} and IL-1 (11). Our results show an impressive upregulation of MIP-1{beta} expression in TBEC upon LPS stimulation. The exact role of this phenomenon with a potential monocyte/macrophage recruitment has to be further elucidated.

In sepsis, TNF-{alpha}, a proinflammatory early response cytokine, is released after exposure to LPS and in turn activates secondary inflammatory cascades consisting of cytokines, lipid mediators, and reactive oxygen species, as well as upregulating cell adhesion molecules, allowing trafficking of inflammatory cells to the site of inflammation (5, 8). Our results suggest that this important cytokine also participates in the inflammatory response of the upper respiratory compartment in vivo. Whether it also has a precursor function for other inflammatory mediators as described previously has to be further explored in detail (18).

ALI is characterized by the accumulation of large numbers of neutrophils from the systemic circulation into the infected lungs. CINC-1 and MIP-2 are two important neutrophil chemoattractants. They have been shown to play an essential role in neutrophil recruitment in acute lung injury (13, 32, 48). As assessed by our chemotaxis assays, LPS-stimulated TBEC also produced neutrophil chemotactic substances, which would implicate an accumulation of neutrophilic effector cells into the respiratory compartment induced by inflammatory mediators, released by LPS-stimulated TBEC.

A further aim of this study was to explore the expression pattern of epithelial cell adhesion molecule in TBEC. These data demonstrated increased production of adhesion molecules in TBEC in vitro upon LPS stimulation. A similar expression pattern of ICAM-1 and VCAM-1 was seen in a human cell line of bronchial epithelial cells upon TNF-{alpha} stimulation (22). Earlier in vivo experiments revealed evidence of increased ICAM-1 expression in bronchial epithelial cells upon LPS stimulation, assuming a certain role of this adhesion molecule in LPS-induced lung injury (5). The present study underlines these previous in vivo findings of enhanced expression of adhesion molecules with in vivo upregulation of VCAM-1 upregulation in epithelial bronchial tissue. Because adhesion molecules are implicated in leukocyte recruitment toward target tissues, the biological relevance of enhanced ICAM-1 and VCAM-1 expression in TBEC was assessed with adherence assays, showing a major role for ICAM-1: two-thirds of neutrophil adhesion was completely abandoned in the presence of ICAM-1 antibodies. The same phenomenon was witnessed for VCAM-1, again with 55% blocking of adhesion using VCAM-1 antibody. Our results imply that, at least in vitro, neutrophil adherence seems to be a selective ICAM-1- and VCAM-1 process. Furthermore, several pieces of evidence obtained in our study argue for an adhesion molecule-dependent mechanism of TBEC killing induced by activated neutrophils. Blocking ICAM-1 or VCAM-1 with respective antibodies resulted in decreased TBEC necrosis. This is a very interesting point, because it would indicate that an antibody treatment could reduce epithelial cell damage in endotoxin-induced epithelial injury.

Our results suggest that activated neutrophils can lead to damage of the integrity of the epithelial barrier not only in the lower respiratory compartment with AEC but also in the upper compartment (41). When neutrophil-induced AEC and TBEC killing are compared, it is apparent that the upper respiratory compartment response occurs before the lower compartment response, which is likely due to TBEC being the first line of defense against pathogens and toxins. An earlier study revealed evidence for increased adhesion of neutrophils to human airway epithelial cells upon stimulation with interferon-{gamma}, TNF-{alpha}, or IL-1 (21). However, exact implications for this phenomenon were not shown.

Further characterization of epithelial cell death in TBEC demonstrated moderately increased apoptosis rates in TBEC upon LPS stimulation, suggesting a direct effect of LPS on bronchial epithelial cell apoptosis. Signaling pathways leading to apoptosis can generally be divided into the extrinsic (surface receptor mediated) and intrinsic (mitochondrial dependent) pathway. The former pathway is triggered after ligand binding to "death" receptors of the TNF superfamily (1). In contrast, the intrinsic pathway is activated without the involvement of death receptors but with alteration of mitochondrial permeability. It was of interest to obtain evidence in LPS-injured TBEC that the mitochondrial pathway might be dominant in the process of apoptosis. Other investigators have postulated a similar apoptotic pathway in bronchial epithelial cells in vivo in murine lung exposed to toxic substances such as 1,1-dichoroethylene (27). Elucidation of these mechanisms may implicate the possibility of novel therapeutic interventions.

The role of apoptosis in the development of ALI remains unclear. Apoptosis might affect the integrity of the bronchial wall with increased permeability and, therefore, disturbance of the main barrier between the environment and respiratory system. It also might be considered as a protective response by the host to pathogens, with the suicide of individual cells enhancing the survival of the multicellular organisms as a whole (53). In type II AEC, extensive apoptosis is largely responsible for the disappearance of these cells in the resolution phase of ALI (2).

Our present results characterize the in vitro inflammatory response in TBEC in the endotoxin-induced lung injury. Improved understanding of the role and contribution of the upper epithelial respiratory compartment to the process of ALI/ARDS will lead to a better understanding of pathophysiological disease processes and the mechanisms that could be affected by novel treatments designed to control such conditions.


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This study was supported by Gebert Ruef Foundation, Switzerland; Jubiläumsstiftung der Schweizerischen Lebensversicherung- und Rentenanstalt; Novartis Stiftung für medizinisch-biologische Forschung, Switzerland; Theodor and Ida Herzog-Egli Stiftung, Switzerland; and Stiftung für wissenschaftliche Forschung, University of Zurich.


   FOOTNOTES
 

Address for reprint requests and other correspondence: B. Beck-Schimmer, Institutes of Anesthesiology and Physiology, Univ. of Zurich Medical School, Winterthurerstrasse 190, CH-8057 Zurich, Switzerland (e-mail: Beatrice_Beck.Schimmer{at}access.unizh.ch)

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

* S. Neff and B. Roth Z'graggen contributed equally to this work. Back


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 Am. J. Physiol. Lung Cell. Mol. Physiol.Home page
P. S. Tang, M. Mura, R. Seth, and M. Liu
Acute lung injury and cell death: how many ways can cells die?
Am J Physiol Lung Cell Mol Physiol, April 1, 2008; 294(4): L632 - L641.
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