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The Journal of Immunology, 2002, 168: 3577-3585.
Copyright © 2002 by The American Association of Immunologists

Activation of Protease-Activated Receptor (PAR)-1, PAR-2, and PAR-4 Stimulates IL-6, IL-8, and Prostaglandin E2 Release from Human Respiratory Epithelial Cells1

Nithiananthan Asokananthan*, Peter T. Graham*, Joshua Fink*, Darryl A. Knight{dagger}, Anthony J. Bakker{ddagger}, Andrew S. McWilliam*, Philip J. Thompson{dagger} and Geoffrey A. Stewart2,*

* Division of Inflammation and Infectious Diseases, Department of Microbiology, {dagger} Asthma & Allergy Research Institute and Department of Medicine, and {ddagger} Department of Physiology, University of Western Australia, Nedlands, Western Australia, Australia


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Epithelia from many tissues express protease-activated receptors (PARs) that play a major role in several different physiological processes. In this study, we examined their capacity to modulate IL-6, IL-8, and PGE2 production in both the A459 and BEAS-2B cell lines and primary human bronchial epithelial cells (HBECs). All three cell types expressed PAR-1, PAR-2, PAR-3, and PAR-4, as judged by RT-PCR and immunocytochemistry. Agonist peptides corresponding to the nascent N termini of PAR-1, PAR-2, and PAR-4 induced the release of cytokines from A549, BEAS-2B, and HBECs with a rank order of potency of PAR-2 > PAR-4 > PAR-1 at 400 µM. PAR-1, PAR-2, and PAR-4 also caused the release of PGE2 from A549 and HBECs. The PAR-3 agonist peptide was inactive in all systems tested. PAR-1, PAR-2, or PAR-4, in combination, caused additive IL-6 release, but only the PAR-1 and PAR-2 combination resulted in an additive IL-8 response. PAR peptide-induced responses were accompanied by changes in intracellular calcium ion concentrations. However, Ca2+ ion shutoff was ~2-fold slower with PAR-4 than with PAR-1 or PAR-2, suggesting differential G protein coupling. Combined, these data suggest an important role for PAR in the modulation of inflammation in the lung.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Asthma is a disease characterized by reversible airway obstruction, bronchial hyperreactivity, and inflammation, and is associated with a predisposition toward atopy. Allergy to a variety of allergens has been described, but allergy to the house dust mite has been shown to be an independent risk factor for the development of disease (1). The precise relationship between allergy and asthma is incompletely understood, but the respiratory epithelium is important because it is the first tissue to meet inhaled allergen and is capable of releasing mediators and cytokines known to be associated with both diseases (reviewed in Ref. 2). For example, respiratory epithelial cells synthesize a variety of proinflammatory cytokines and chemokines such as IL-1, IL-6, GM-CSF, IL-8, macrophage-inflammatory protein, eotaxin, and RANTES (3, 4, 5, 6), as well as anti-inflammatory mediators such as PGE2 and NO (reviewed in Ref. 7), all of which regulate inflammation through their effects on cell recruitment, activation, and survival.

A variety of agents has been shown to stimulate cytokine and mediator release from airway epithelium, including eosinophil granule proteins, fungi, viruses, and bacteria (8, 9, 10, 11). In addition, we and others have also shown that endogenous and exogenous peptidases, including allergens, are also significant modulators of epithelial function. For example, neutrophil elastase and allergenic cysteine and serine peptidases from house dust mites and fungi have been shown to stimulate IL-6 and IL-8 release (12, 13, 14), although how such peptidase-induced effects were mediated at the cellular level was not described. More recently, the involvement of members of a recently identified G protein-coupled family of cell surface receptors designated protease-activated receptors (PARs)3 has been implicated (13, 15, 16).

To date, four distinct PARs have been described (17, 18, 19, 20): PAR-1, PAR-2, and PAR-3 are encoded by genes located on chromosome 5q13, whereas the PAR-4 gene is located on chromosome 19q12 (21, 22, 23, 24). Each receptor has been shown to modulate a variety of physiological processes such as cytokine and mediator release, vasodilation, platelet aggregation, cellular proliferation, and smooth muscle contraction or relaxation (reviewed in Refs. 25, 26), all of which may be relevant in the pathogenesis of allergic disease. Thrombin activates PAR-1 and PAR-3 (17, 21), whereas trypsin and mast cell tryptase activate PAR-2 (27). In contrast, PAR-4 is activated by thrombin and trypsin as well as cathepsin G (20, 28). These peptidases activate PAR via cleavage of the extracellular N-terminal domain, which then enables the new N terminus (now referred to as a tethered ligand) to interact distally within the same molecule to activate G protein-coupled signal transduction pathways (29). These receptors can also be activated without proteolytic cleavage using five to six residue peptides corresponding to the new amino termini of the cleaved receptors (17).

Although the distribution of some of the PARs has been studied in several tissues, the precise locations and physiological roles of all four PARs in the human respiratory epithelium function are unclear. Similarly, detailed knowledge of the types of exogenous and endogenous peptidases present in the lung capable of activating them is limited. In the present study, we performed experiments to determine whether all members of the PAR family were expressed on respiratory epithelial cells and whether activation of individual PAR caused the production of the cytokines IL-6 and IL-8 as well as the prostanoid PGE2, since each is likely to be important in both asthma and atopy per se. Similarly, we performed experiments using thrombin and trypsin, given that both have been directly or indirectly implicated in allergic disease (30) or are known to be located in the lung (31).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials

Synthetic, agonist peptides, as well as control PAR peptides were synthesized with amidated C termini (purity >85%; Protein Facility, University of Western Australia, Perth, Australia). The sequences of the active and control peptides, respectively, were: PAR-1, SFLLRN-NH2 and FSLLRN-NH2; PAR-2, SLIGKV-NH2 and LSIGKV-NH2; PAR-3, TFRGAP-NH2 and FTRGAP-NH2; and PAR-4, GYPGQV-NH2 and GYPGVQ-NH2. Given that the human PAR-1 agonist peptide at high concentration has been shown to activate PAR-2 (32), the more specific frog PAR-1 agonist peptide TFLLRN-NH2 (control peptide, FTLLRN-NH2) was also examined. Thrombin was purchased from CSL (Melbourne, Australia) and Sigma-Aldrich (St. Louis, MO). Tissue culture reagents and molecular biology reagents were purchased from Life Technologies (Melbourne, Australia) and Stratagene (La Jolla, CA), respectively. General chemicals were purchased from either BDH (Kilsyth, Victoria, Australia) or Sigma-Aldrich, unless otherwise stated. The human pulmonary epithelial type II cell line A549 and the SV40-transformed human bronchial epithelial cell line BEAS-2B were obtained either from the American Type Culture Collection (Manassas, VA) or C. Harris (National Institutes of Health, Bethesda, MD), respectively. Primary human bronchial epithelial cells (HBECs) were purchased from Clonetics (San Diego, CA) or generated, as described below.

Protease activity determinations

Trypsin activity was expressed in molar terms after determining the percentage of active enzyme using the active site titrant, p-nitrophenyl p'-guanidino benzoate, as described previously (33), whereas thrombin activity was expressed in U ml-1, as determined by the manufacturer. However, the protease activity of thrombin was determined using N-benzoyl-DL-arginine p-nitroanilide, as described previously (34), to confirm peptidase activity before experimentation. The thrombin supplied by CSL (0.5 nM p-nitroaniline released min-1 mg-1) was found to be ~50-fold less active on this basis than that supplied by Sigma-Aldrich (23.4 nM p-nitroaniline released min-1 mg-1). Thrombin from the former source was used in cell culture experiments, whereas that from the latter was used in Ca2+ flux studies.

Cell culture

The A549 cells were cultured as described previously (35). For initial growth, cells were seeded into 75-cm2 tissue culture flasks (Nunc, Naperville, IL) and grown to confluence in Ham’s F12 Kaighn’s Modification (Life Technologies) medium supplemented with 10% (v/v) FCS and antibiotics. For subsequent experiments, cells from the flasks were trypsinized and seeded into 24-well tissue culture plates at a density of 5 x 104 cells/well-1 and grown to 80% confluency. At this time, the cells were then incubated in basal medium for an additional 24 h. The BEAS-2B cells were grown under conditions similar to those described previously (13). In order for the BEAS-2B cells to adhere to the tissue culture plastic, flasks and plates were coated with a solution of 30 µg ml-1 rat collagen S, 1 mg ml-1 human fibronectin, and 100 µg ml-1 BSA in Lechner and La Veck medium (LHC) basal medium (Biofluids, Rockville, MD) for 6 h at 37°C in 5% (v/v) CO2. The flasks and plates were then washed with serum-free medium comprising LHC basal medium supplemented with insulin, hydrocortisone, epidermal growth factor, bovine hypothalamus extract, cholera toxin, transferrin, and retinol acetate, as previously described (36). Whenever BEAS-2B or A549 cells were needed for experiments, mediumwas changed every 3 days until cells were confluent. Confluent cultures from the flasks were subsequently seeded onto 24-well plates or slides, as appropriate. At confluence, the medium was replaced with LHC basal medium.

HBECs either were derived from human bronchi, as described previously (37), or commercially obtained from Clonetics. For in-house derived cultures, bronchial tissue pieces, free of all visible blood vessels, were stripped from the airway wall and cut into 2-mm2 pieces that were then oriented, epithelial surface uppermost, on plastic culture dishes. The tissue explants were allowed to adhere onto rat collagen S-coated (30 µg ml-1) plastic culture dishes at 37°C for 16 h, after which growth factor-supplemented LHC-9 medium was added gently to avoid dislodging the tissue. The medium was changed at 2- to 3-day intervals until growth had reached the edge of the dish (12–20 days). The explants were removed, and cells were cultured for an additional 24 h before use in medium devoid of supplements. Commercially obtained HBECs were seeded into plastic flasks and grown in bronchial epithelial cell growth medium (Clonetics) supplemented with bovine pituitary extract (52 mg ml-1), hydrocortisone (0.5 mg ml-1), human recombinant epidermal growth factor (0.5 mg ml-1), epinephrine (0.5 mg ml-1), transferrin (10 mg ml-1), insulin (5 mg ml-1), retinoic acid (0.1 mg ml-1), triiodothyronine (6.5 mg ml-1), gentamicin (50 mg ml-1), and amphotericin B (50 mg ml-1). Medium was changed every 48 h until cells were 90% confluent. Cells were then passaged and seeded onto 24-well plates and, at confluence, medium was replaced with bronchial epithelial cell growth basal medium. Cells were used between days 7 and 14 postseeding. At all stages of culture, cells were maintained at 37°C in 5% (v/v) CO2.

Stimulation of epithelial cells

Epithelial cells were grown in appropriate serum-free basal media for a total of 24 h and exposed to thrombin (0.05–500 U ml-1), trypsin (0.0125–1.25 nM), or PAR peptides (50–500 µM). In experiments designed to study PAR-induced cytokine and mediator secretion at different times during culture, cells were exposed to the optimum concentration of peptide for 24 h. Culture supernatants were collected, centrifuged at 12,000 x g for 5 min at 4°C, and stored frozen. At the conclusion of each experiment, cells were detached, and viability and cell number were determined by trypan blue exclusion. Epithelial cells exposed to the various additives were analyzed for potential cytotoxic reactions by measuring the release of lactate dehydrogenase using a spectrophotometric assay (38).

Immunohistochemistry

Immunolocalization of PARs on epithelial cells was performed using cells cultured in eight-well chamber slides (LAB-Tek II; Nunc) to 80% confluency. The cells were washed in PBS and fixed in 4% (v/v) paraformaldehyde in PBS. Endogenous peroxidase activity was quenched by incubating tissue sections in 3% (v/v) H2O2 for 5 min, and nonspecific binding was blocked by the addition of three drops of DAKO Biotin Block (DAKO, Carpinteria, CA). Cells were then incubated with Abs raised against individual PARs. These included: anti-PAR-1 (WEDE 15), a mouse monoclonal raised against 51KYEPFWEDEEKNES64 (39); anti-PAR-2 (SAMII), a mouse monoclonal raised against 37SLIGKVDGTSHVTG50 (40); anti-PAR-3, a rabbit Ab raised against 37TLPIKTFRGAPPNSFEEFP55; and anti-PAR-4, rabbit Ab raised against 28EDDSTPSLLPAPRGYPGQV39. The mouse mAbs were used as ascites fluid, and rabbit Abs were used after affinity purification. The rabbit Abs were prepared commercially (Chiron, Clayton, Australia). The cells were incubated with either the primary Ab or isotype-matched primary Ab control in 4% (v/v) normal goat serum. Following two washes in PBS, sections were incubated with biotinylated secondary Abs, followed by incubation with S-HRP, and visualized by diaminobenzidine, as described previously (41).

RT-PCR

Total RNA was prepared from the primary epithelial cells and cell lines using Tri-Reagent (Molecular Research Center, Cincinnati, OH), as described by the manufacturer. Briefly, cells were lysed and the homogenates were transferred to 1.5-ml microcentrifuge tubes. A total of 200 µl chloroform was added, and each tube was shaken by hand and left at room temperature for 10 min. Tubes were then centrifuged in a microfuge at 13,000 rpm for 15 min at room temperature, after which the aqueous phase was transferred to new tubes containing 0.5 ml of isopropanol and shaken. The tubes were incubated at room temperature for 10 min and then centrifuged at 13,000 rpm for 8 min. The supernatants in each tube were discarded, and 1 ml 75% (v/v) ethanol was added to the pelleted RNA. The pellets were resuspended, and tubes were centrifuged at 13,000 rpm for 5 min, after which the supernatants were discarded and the pellets were air dried and dissolved in 30 µl of diethyl pyrocarbonate-treated water. RNA was quantitated spectrophotometrically, and integrity was analyzed by electrophoresis on a 5% (v/v) formaldehyde, 1% (w/v) agarose gel. cDNA was prepared by reverse transcriptase using a commercial RT-PCR kit, and reactions were performed according to the manufacturer’s instructions. Briefly, 10 µg of RNA was added to a 50-µl reaction volume containing 3 µl of random primers, 5 µl of 10x first-strand buffer, 1 µl of RNase Block (40 U/µl), 2 µl of 100 mM dNTPs, and 1 µl of Moloney murine leukemia virus reverse transcriptase (50 U µl-1). The reaction mix was incubated at 37°C for 60 min, then at 90°C for 5 min. Forward and reverse primers used for amplifying human PARs were prepared commercially based on published sequence data (29): PAR-1, sense 5'-TGTGAACTGATCATGTTTATG-3', antisense 5'-TTCGTAAGATAAGAGATATGT-3' (PCR product, 708 bp); PAR-2, sense 5'-AGAAGCCTTATTGGTAAGGTT-3', antisense 5'-AACATCATGACAGGTCGTGAT-3' (PCR product, 582 bp); PAR-3, sense 5'-CTGATACCTGCCATCTACCTCC-3, antisense 5'-AGAAAACTGTTGCCCACACC-3' (PCR product, 382 bp); PAR-4, sense 5'-ATTACTCGGACCCGAGCC-3', antisense 5'-TGTAAGGCCCACCCTTCTC-3' (PCR product, 392 bp). Amplification of {beta}-actin, with the sense and antisense primer pair 5'-GGCTCTTCCAGCCTTCCTTCCT-3' and 5'-CACAGAGTACTTGCGCTCAGGAGG-3' (PCR product 240 bp), acted as an internal control. The conditions for amplification were: PAR-1, PAR-2, 94°C for 45 s for 1 cycle, 55°C for 45 s for 35 cycles, 72°C for 2 min and 30 s for 35 cycles, and 72°C for 10 min for 1 cycle; PAR-3, PAR-4, and {beta}-actin, 94°C for 45 s for 1 cycle, 65°C for 45 s for 1 cycle, 72°C for 2 min and 30 s for 35 cycles, and 72°C for 10 min for 1 cycle. Each PCR was performed in a 50-µl reaction volume containing 0.25 µl of cDNA, 0.125 µl of 100 mM dNTPs, 2.5 µl of 10x buffer for cloned PFu polymerase, and 0.2 µl of cloned PFU polymerase water to 25 µl. Electrophoresis was conducted on 2% (w/v) analytical grade agarose gels that were subsequently stained with ethidium bromide. PCR products were purified using a QIAquick gel extraction kit (Qiagen, Melbourne, Australia), and their identities were confirmed using an Applied Biosystems (Foster City, CA) ABI 377 automated DNA sequencer (Department of Immunology, Royal Perth Hospital, Perth, Western Australia).

Determination of IL-1{beta}, IL-6, IL-8, and TNF-{alpha} production

IL-1{beta}, IL-6, and IL-8 production was determined using specific ELISA, as described previously (13). Briefly, 96-well plates (Maxisorp; Nunc) were coated with 100 µl/well of the appropriate Ab (250 ng ml-1 in 0.1 M NaHCO3/NaCO3 buffer, pH 9.6) and incubated overnight at 4°C. The plates were then washed three times with washing buffer (PBS, pH 7.5, containing 0.5% (v/v) Tween 20) and blocked by incubating with 100 µl of blocking buffer/well (washing buffer containing 1% (w/v) BSA) at room temperature for 1 h. The plates were washed three times with washing buffer before 100 µl of test or standards were added to the wells. Plates were incubated overnight at 4°C, washed, and incubated with biotinylated secondary Ab (100 µl of a 250 ng ml-1 stock in blocking buffer/well) at room temperature for 1 h. After washing, 100 µl of peroxidase-labeled streptavidin (125 ng ml-1; Kirkegaard & Perry Laboratories, Gaithersburg, MD) was added to each well, incubated at room temperature for 30 min, and, after washing, incubated with 100 µl of peroxidase substrate (K-Blue ELISA Substrate; Graphic Scientific, Brisbane, Australia) per well. Reactions were terminated by the addition of 100 µl of 1 M phosphoric acid/well, and OD was determined using a microplate reader (Spectramax 250; Molecular Devices, Menlo Park, CA) at 450 nm. The concentrations of cytokines in each sample were determined by interpolation from the standard curve using the SoftMax-Pro software (Molecular Devices) and expressed as picograms of 2.5 x 105 cells-1. Second Abs used in ELISA were obtained from BD PharMingen (San Diego, CA). TNF-{alpha} was measured using the L-929 bioassay, as described previously (42). Briefly, L-929 cells were cultured in 96-well microtiter plates at 5 x 104 cells/well-1 in RPMI 1640 containing 10% (v/v) FCS for 20 h. Medium was then replaced with 50 µl of medium containing 4 µg/ml actinomycin D (Boehringer Mannheim, Castle Hill, Australia). Standards and samples (50 µl) were added to each well in duplicate, and the cells were incubated for an additional 20 h, after which the medium was removed and the cells were stained with 0.1% (w/v) crystal violet in 1% (v/v) acetic acid for 15 min to detect lysis. Plates were then washed three times with distilled water and allowed to dry; remaining cells were solubilized with 100 µl of 1% (w/v) SDS in water; and OD was measured at 590 nm.

Detection of PGE2

PGE2 was measured using a competitive enzyme immunoassay (Cayman Chemical, Ann Arbor, MI), as described previously (35). Briefly, 96-well plates precoated with the capture Ab (goat anti-mouse Ab) were loaded with 50 µl of samples or standards and incubated with 50 µl of PGE2 tracer and 50 µl of PGE2 mAb overnight at 4°C. After three washes in wash buffer, 200 µl of Ellman’s reagent was added to the plate and allowed to incubate for 1 h for the color to develop. OD at 405 nm was determined, and PGE2 production was expressed as picograms of 2.5 x 105 viable cells-1.

Measurement of cytosolic-free calcium (Ca2+)

Changes in cytosolic Ca2+ concentration were measured using the fluorescent Ca2+ indicator, indomethacin (Indo)-1 acetoxymethylester (AM) (Teflabs, Austin, TX). The ratio of fluorescence emission at 405 and 490 nm was measured using a spectrophotometer (Cairn, Faversham, Kent, U.K.) attached to an inverted microscope (Nikon, Tokyo, Japan), configured for epifluorescence. The excitation wavelength of 340 nm was provided by a variable monochromator system (Cairn). A549 cells were grown on a coverslip to 80% confluency and loaded with Indo-1 by incubation in RPMI 1640 (medium free of Phenol Red) containing 6 µM membrane-permeable form of the Ca2+ indicator, Indo-1 AM, and 0.1% (w/v) Pluronic F-127 (Molecular Probes, Eugene, OR) for 45 min at room temperature. After incubation, cells were washed with two changes of RPMI 1640 to remove the excess dye and then exposed to the various test substances after establishing baseline. The ratio of the emission intensities was used as a measure of changes in cytosolic Ca2+. Calcium flux shutoff was determined as the time taken from agonist addition to half-maximum response, as described previously (43). In experiments designed to measure receptor specificity, A549 cells loaded with Indo-1 dye were initially exposed to either peptidases or peptides, followed by the second stimulus, after ensuring that the calcium flux reached a baseline, or after 3 min if no changes in Ca2+ were observed.

Statistical analyses

Unless stated otherwise, data are expressed as mean ± SEM. Statistical significance between means was determined using ANOVA or the Student’s t test using Microsoft Excel for the Macintosh. The Bonferoni correction was used for multiple t test comparisons.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Thrombin and trypsin induce cytokine release from A549 cells

Thrombin and trypsin caused significant release of IL-6 and IL-8 from A549 cells in a dose-dependent manner over a 24-h period (Fig. 1Go) when compared with medium controls. The mean maximum concentrations of IL-6 released in response to thrombin and trypsin were obtained with 500 U and 0.125 nM, respectively (Fig. 1Go, left panel). The mean maximum concentrations of IL-8 released on exposure to thrombin and trypsin were obtained with 50 U and 0.0125 nM, respectively (Fig. 1Go, right panel). Apart from thrombin-induced IL-6 production, IL-6 and IL-8 production was reduced at peptidase concentrations higher than those shown to be optimal (Fig. 1Go).



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  		FIGURE 1. Endogenous peptidases promote cytokine release from the human respiratory epithelial cell line A549 in a dose-dependent manner. Cell monolayers were cultured for 24 h with basal medium devoid of serum, then stimulated with increasing concentrations of either thrombin, trypsin, or medium alone (Control). The presence of IL-6 and IL-8 in supernatants was measured by ELISA, and data were expressed as mean ± SEM 105 cells-1 from three independent experiments performed in quadruplicate. Significant differences in mean peptidase-induced cytokine responses compared with medium (Control) were determined using the Student’s t test and Bonferoni’s correction; *, p < 0.05; **, p < 0.001.

 
Respiratory epithelial cells express PARs

Before assessing the effects of PAR agonist peptides on respiratory epithelial cell function, the presence of individual PAR on A549 and BEAS-2B cell line, as well as HBECs, was determined by immunocytochemistry (Fig. 2Go) and RT-PCR (Fig. 3Go). All four PARs were detected by both techniques. The PAR amplicons obtained in the RT-PCR experiments were sequenced and found to correspond to published sequences (data not shown).



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  		FIGURE 2. Immunohistochemical analysis of PAR on respiratory epithelial cell lines. A549 (A–E) and BEAS-2B (F–J) lung epithelial cell lines were stained using an immunoperoxidase method.

 


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  		FIGURE 3. RT-PCR analysis of PAR expression by HBECs (A), A549 (B), and BEAS-2B (C) epithelial cell lines. RNA was extracted, reverse transcribed, and amplified with specific primers for PARs and {beta}-actin. Products were separated on 2% (w/v) wide-range agarose gels, stained with ethidium bromide, and photographed. Lane 1, 100-bp DNA ladder; lanes 2–6, PAR-1 (708 bp), PAR-2 (582 bp), PAR-3 (382 bp), PAR-4 (392 bp), and {beta}-actin (240 bp), respectively.

 
Respiratory epithelial cell lines and HBECs secrete cytokines in response to PAR agonist peptides

Initial experiments were performed on the A549 cell line, and the results obtained showed that PAR-1, PAR-2, and PAR-4 agonist peptides stimulated the secretion of both IL-6 and IL-8, with a rank order of potency of PAR-2 > PAR-4 > PAR-1 at optimal concentrations for each of 400 µM (data not shown). However, as human PAR-1 may activate PAR-2 (32), the experiments were repeated using frog PAR-1 agonist peptide. A similar potency ranking and optimal concentrations were obtained (Fig. 4Go, top left). PAR-3 agonist peptide did not stimulate cytokine secretion at any concentration or time point tested. Neither IL-1{beta} nor TNF-{alpha} release was detected in response to any PAR agonist stimulation (data not shown).



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  		FIGURE 4. PAR-1, PAR-2, and PAR-4 agonist peptides promote cytokine release from the A549 cells in a dose- and time-dependent manner. Cell monolayers were cultured for 24 h with basal medium devoid of serum, then stimulated with increasing concentrations of PAR agonist peptides (top two panels, frog PAR-1, {square}; PAR-2, •; PAR-4, {blacksquare}) for 24 h. Bottom two panels, Cell monolayers were cultured for 24 h with basal medium devoid of serum, then stimulated with agonist peptides at 400 µM (frog PAR-1, {square}; PAR-2, •; PAR-4, {blacksquare}; medium, {circ}), and cytokine release was determined by harvesting the supernatants at 3, 6, 9, 12, 18, and 24 h. Data are expressed as mean ± SEM 2.5 x 105 cells-1 from three independent experiments performed in quadruplicate. The concentrations of IL-8 secreted in response to frog PAR-1 at 3, 6, and 12 h were not significantly different from the medium-induced responses at the corresponding time points, as determined using ANOVA.

 
Similar experiments were performed using the BEAS-2B cell lines and HBECs Again, qualitatively similar results to those obtained with A549 cells were noted (Fig. 5Go, top panels), although quantitative differences were observed. For example, IL-6 production in response to PAR peptides was similar in A549 and BEAS-2B cell lines with regard to mean concentration of cytokine produced, although constitutive cytokine production was higher with the BEAS-2B cell line (Fig. 5Go, middle panels). In contrast, PAR peptide-induced IL-6 production from the HBECs was approximately twice that observed with A549 and BEAS-2B cell lines, and constitutive levels were relatively higher in both HBECs and BEAS-2B than the A549 cell line. PAR peptide-induced IL-8 responses in the A549 cell line and HBECs were approximately twice that obtained with the BEAS-2B cell line. Control peptides had little effect on cytokine secretion from either cell line or HBECs (Fig. 5Go, bottom panels). Examination of medium from cells exposed to either peptidase or peptides for lactate dehydrogenase indicated that PAR treatment did not result in cytotoxic reaction (data not shown).



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  		FIGURE 5. PAR agonist peptides promote cytokine release from A549, BEAS-2B cells, and HBECs. Cell monolayers were cultured with basal medium devoid of serum or growth factors for 24 h, then stimulated with both agonist (+) and control peptides (-) for 24 h. Data are expressed as mean ± SEM 2.5 x 105 cells-1 from three independent experiments performed in quadruplicate. Significant differences in means between PAR agonist peptide-induced and control peptide-induced (Control) responses at 400 µM concentration were determined using Student’s t test (*, p < 0.05); PAR-2 agonist peptide-induced responses were significantly different from that obtained with PAR-1 or PAR-4 peptide with all three cell types ({dagger}, p < 0.05-p < 0.005). PAR-4 agonist peptide-induced responses were significantly different from PAR-1 agonist peptide-induced cytokine responses ({dagger}{dagger}, p < 0.05) with all three cell types.

 
Respiratory epithelial cell lines respond to combinations of PAR agonist peptides

Exposure of the A549 cells to PAR-1 and PAR-2 agonist peptides in combination (total final concentration, 800 µM) resulted in an additive IL-6 response (Fig. 6Go, left panel). A similar effect was seen with either PAR-1 or PAR-2 in combination with PAR-4. In contrast, additive effects on IL-8 release were observed only with the PAR-1 and PAR-2 combination (Fig. 6Go, right panel). Responses obtained with individual peptides at 800 µM were not significantly different from those obtained at 400 µM (Fig. 6Go). Exposure of the cells to PAR-3 agonist peptide in any combination did not influence the release of either cytokine.



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  		FIGURE 6. Additive effect of PAR agonist peptides on the secretion of cytokines from A549 cells. Cell monolayers were cultured with basal medium without serum for 24 h before stimulation, and then treated with agonist peptide (+) or control peptide (-) either alone or in combination with another PAR peptide. Data are expressed as mean ± SEM 2.5 x 105 cells-1 from three independent experiments performed in quadruplicate. Significant differences in mean cytokine production between combined PAR agonist peptides compared with individual PAR peptide-induced responses at 800 µM concentration were determined using Student’s t test; *, p < 0.05; **, p < 0.005.

 
Respiratory epithelial cell lines and HBECs release PGE2 in response to PAR agonist peptides

Exposure of the A549 cells and HBECs to PAR agonist peptides showed that PAR-1, PAR-2, and PAR-4 peptides also induced the production of PGE2 (Fig. 7Go), although both constitutive and PAR-induced PGE2 production were ~10-fold higher in the HBECs (Fig. 7Go, left panel). To determine whether the PAR-induced cytokine responses described above were dependent on this PGE2 production, A549 cells were incubated in the presence of agonist peptide and Indo. However, preincubation of cells with Indo before PAR-1 and PAR-2 peptide exposure did not significantly alter the production of either IL-6 or IL-8 when compared with that obtained with cells exposed to peptides alone (Fig. 8Go).



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  		FIGURE 7. PAR-1, PAR-2, and PAR-4 peptides promote PGE2 release from A549 cells and HBECs. Cell monolayers were cultured with basal medium without serum for 24 h before stimulation and then treated with either agonist (+) or control (-) PAR peptides (400 µM) for 24 h. The concentrations of PGE2 in supernatants were determined by ELISA. Data are expressed as mean ± SEM 2.5 x 105 cells-1 from three independent experiments performed in quadruplicate. Significant differences in means between PAR agonist peptide-induced and control peptide-induced responses were determined using Student’s t test; *, p < 0.05; **, p < 0.005.

 


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  		FIGURE 8. Indo treatment does not ablate PAR-induced cytokine release from A549 cells. Cell monolayers were cultured with basal medium without serum for 24 h before stimulation. The cells were then pretreated with Indo in DMSO before the exposure to 400 µM of either PAR-1 or PAR-2 agonist peptides for 24 h. Data are expressed as mean ± SEM 2.5 x 105 cells-1 from experiments performed in triplicate. Statistically significant differences in mean PAR peptide-induced cytokine production in the presence or absence of Indo were determined using ANOVA; NS, no significant difference.

 
Changes in intracellular calcium induced by peptidase and PAR peptides

To examine the association between intracellular calcium levels and PAR-mediated cytokine production, the changes in free intracellular calcium were monitored. Optimal concentration of both thrombin (10 U ml-1) and trypsin (0.125 nM) induced changes in the intracellular Ca2+ of the A549 cells (Fig. 9Go, top panels) within a few seconds, as judged by the fluorescence ratio. Similarly, the PAR-1, PAR-2, and PAR-4 agonist peptides, at concentrations ranging from 50 to 400 µM, also significantly and abruptly caused an increase in intracellular Ca2+ (Fig. 9Go, bottom panels). The optimum concentrations of PAR-1 and PAR-2 agonist peptides were both shown to be 100 µM, whereas that for PAR-4 peptide was shown to be 300 µM. In this regard, PAR-4 agonist peptide did not always produce a Ca2+ response greater than that achieved with the PAR-4 control peptide. Neither PAR-3 agonist peptide nor PAR-1, PAR-2, and PAR-4 control peptides induced any significant increase in the mean fluorescence ratio over time, although PAR-3 control peptide induced a small, slow, but significant increase in the mean resting fluorescence ratio (Table IGo). The reason for this response is unclear, but is unlikely to be due to the effects of bleaching of the Ca2+ indicator, as addition of medium alone did not produce similar changes. The time taken to shut off peptide-induced calcium responses was measured, and the data obtained showed that it was much slower in PAR-4-activated cells compared with either PAR-1, PAR-2, thrombin, or trypsin (Table IGo). Similarly, it was significantly slower with trypsin-treated cells compared with thrombin-treated cells. However, frog PAR-1 agonist peptide and thrombin had approximately similar shut off times.



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  		FIGURE 9. PAR agonist peptide-induced cytosolic Ca2+ responses in A549 cells. A549 cells were loaded with Indo-1AM and incubated with thrombin (Thr, 25 U ml-1) and trypsin (Try, 0.25 nM) or incubated with 400 µM of either agonist peptide (Pep) or control peptide (Con). Responses were measured over 400 s and expressed as a fluorescence ratio (405/485 nm).

 

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  		Table I. Kinetics of PAR agonist and peptidase-induced Ca2+ flux in A549 cells

 
Specificity of the PAR-1 and PAR-2 receptors on respiratory epithelial cells

The effect of exposing A549 cells sequentially to thrombin, trypsin, PAR-2, frog PAR-1, or human PAR-1 peptides was determined. Exposure of the A549 cells to thrombin before either frog PAR-1 (Fig. 10GoA) or human PAR-1 agonist peptide (data not shown) treatment resulted in desensitization to the latter, but not to subsequent treatment with PAR-2 peptide (Fig. 10GoB) or trypsin (Fig. 10GoC). Exposure of cells to trypsin before frog PAR-1 (Fig. 10GoD) or human PAR-1 (data not shown) and human PAR-2 agonist peptide (Fig. 10GoE) resulted in desensitization with the latter only. Desensitization was not obtained when cells were treated with frog PAR-1 agonist peptide and then PAR-2 peptide (Fig. 10GoF). However, when cells were exposed to human PAR-1 and then treated with human PAR-2, desensitization was observed (Fig. 10GoG). Desensitization was not observed when this treatment process was reversed (Fig. 10GoH).



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  		FIGURE 10. Desensitization of PARs in A549 cells. A549 cells were loaded with Indo-1AM and incubated with either peptidase or PAR peptide. A549 cells were incubated with thrombin (20 U ml-1) or trypsin (0.25 nM), followed by incubation with 100 µM of either frog PAR-1, human PAR-2 agonist peptides, or trypsin (0.25 nM) (A–E). A549 cells were also incubated with frog PAR-1 followed by human PAR-2 (F). A549 cells were also incubated with human PAR-1 followed by PAR-2 (100 µM; G), or PAR-2 (100 µM) followed by human PAR-1 (100 µM; H). Arrows indicate addition of second stimulus. Responses were measured over 400 s and expressed as a fluorescence ratio (405/485 nm).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In previous studies, we demonstrated that the endogenous peptidases trypsin and thrombin, as well as the exogenous allergenic peptidases derived from the house dust mite, namely, Der p 1 and Der p 9, triggered the synthesis and release of IL-6 and IL-8 from both primary cultures of human bronchial epithelium and the respiratory epithelial cell line BEAS-2B (13). Although the mechanism(s) underlying these findings was unclear, the available literature on the PAR family led us to believe that they may have been involved (13). Therefore, in the current study, we determined the presence and functionality of PAR in human lung epithelial cells, before assessing their role in allergen-induced events. To this end, we demonstrated that not only were the four known PARs on primary cultures of respiratory epithelial cells and epithelial cell lines, but also that the activation of PAR-1, PAR-2, and PAR-4, but not PAR-3, resulted in cytokine and mediator release.

Initial studies were performed to determine whether known peptidase modulators of PAR function influenced the release of IL-6 and IL-8 from A549 cells. The results from these studies demonstrated that both thrombin (activator of PAR-1, PAR-3, and PAR-4) and trypsin (activator of PAR-2 and PAR-4) induced both time- and concentration-dependent cytokine release. These data are consistent with those reported from our laboratory (13, 44) and elsewhere (45, 46). In the present study, we showed that exposure of cells to increasing concentrations of peptidases resulted in diminished cytokine release, which, on the basis of previous data (13, 45), suggests susceptibility of individual cytokines to proteolysis. This susceptibility may also account for the variation in mean maximum IL-8 concentrations obtained in response to thrombin and trypsin or, alternatively, reflects the involvement of one or more PAR member in cytokine induction (see below). Although the precise role of thrombin and trypsin in asthma is unclear, both thrombin and the trypsin-related mast cell peptidase, tryptase, have been shown to be elevated in sputum from asthmatic patients compared with controls and, in the case of tryptase, enzyme concentration correlated with disease severity (30, 47). Similarly, protease inhibitors have been shown to modulate bronchial activity, suggesting that proteases per se are important in the pathogenesis of this disease (48).

In this study, all four PARs were detected on human primary epithelial cells as well as the commonly used epithelial cell lines, as judged by both RT-PCR and immunohistochemistry. These data confirm and extend those observed in previous studies from our group and elsewhere (49, 50, 51, 52). We showed that both type of cells released IL-6 and IL-8, but not IL-{beta} and TNF-{alpha} when treated with PAR-1, PAR-2, and PAR-4, but not PAR-3 agonist peptides, with a rank order of potency of PAR-2 > PAR-4 > PAR-1. Cytokine responses were not observed with control peptides produced by reversing the position of the first two amino acids in PAR-1 and PAR-2, but not PAR-4 sequence. When this procedure was adopted for the PAR-4 peptide, similar cytokine responses were obtained for both agonist and control peptides (data not shown). This observation is consistent with data obtained using the mouse homologue in an isolated tracheal tissue model (52), and with other studies indicating a preference for residues with a small aliphatic side chain at position 1 in the agonist peptide (53). However, this finding is not consistent with the suggestion that an aromatic side chain is required at position 2 in the agonist peptide (53). When the two C-terminal residues were reversed, the control peptide was rendered inactive, suggesting that this region of the peptide is particularly important in regulating activity.

In other studies, it has been shown that proinflammatory cytokines such as TNF-{alpha}, IL-1{beta}, and the prostanoid PGE2 (54, 55) may also stimulate cytokine release. Thus, it was possible that the PAR-mediated activities observed represented secondary phenomena. In this regard, we failed to detect production of either IL-1{beta} or TNF-{alpha}, although PGE2 production in response to stimulation with PAR-1, PAR-2, and PAR-4 was detected. These findings are consistent with previous findings from our laboratory and elsewhere (52, 56). For example, we showed that mouse PAR-1, PAR-2, and PAR-4 agonist peptides induced epithelium-dependent PGE2 production by isolated murine tracheal sections, which was inhibited by Indo treatment (56). To investigate whether PGE2 might be acting in a paracrine fashion, cells were pretreated with Indo before PAR stimulation. However, treatment with this inhibitor failed to ablate cytokine production in response to PAR-1, PAR-2, or PAR-4, suggesting that the effects observed were primarily associated with PAR activation or with some other factor yet to be determined.

The capacity of PAR peptides to stimulate cytokine release from different cell types has been described previously, particularly for PAR-1 and PAR-2. For example, lung and gingival fibroblasts, keratinocytes, endothelial cells, and intestinal epithelial cells have been shown to produce IL-6, IL-8, and GM-CSF in response to PAR-1 and PAR-2 (46, 57, 58). We and others now show that human respiratory epithelial cells also produce cytokines in response to PAR agonist peptides. In this regard, PAR-2 peptide has been shown to stimulate GM-CSF production from the lung epithelial cells, and we have extended these observations to include IL-6, IL-8, and PGE2. In addition, we have shown that PAR-1 and PAR-4 also induce cytokine release, and that PAR-1, PAR-2, and PAR-4 peptides induce PGE2 responses from human respiratory epithelial cells. With regard to PAR-1, it was possible that these effects were initiated by the activation of either PAR-1, PAR-2, or both based on previously reported data using variety of assays (32, 59, 60). In this regard, our data obtained with the frog PAR-1-activating peptide suggested that this receptor is functional in respiratory epithelium. We also showed an additive effect when combinations of PAR agonist peptides were used to stimulate cells, again suggesting distinct receptor interactions and/or signal transduction mechanisms consistent with previous reports linking PAR-1 activation with coupling of receptors to both Gi and Gq and G12/13 subfamilies, and PAR-4 with Gq, but not Gi (53). These data imply potential enhancement of epithelial responses in the presence of multiple peptidases likely to arise during inflammatory responses.

Activation of PARs by enzymatic cleavage of the extracellular N-terminal region has been shown to result in inositol phospholipid hydrolysis and increased cytosolic-free Ca2+, leading to cellular activation and cytokine release (58). These properties are often used to explore PAR-associated transduction processes, and, in this regard, we showed both qualitative and quantitative differences in PAR-1-, PAR-2-, and PAR-4-induced changes in cytosolic Ca2+. We showed that the magnitude of the fluorescent ratios obtained with PAR-1 and PAR-2 was greater than those seen with PAR-4 in the A549 cell line, and the time taken for shutoff was ~2-fold greater with PAR-4 than with either PAR-1 or PAR-2. This difference in shutoff is consistent with data obtained with other cell types in both native and transfected cells (43, 61), in which sustained responses are thought to reflect the mobilization of Ca2+ from the extracellular space as well as continued receptor occupancy, resulting in a slower uncoupling from G protein-coupled signaling events (62). The physiological significance of differential Ca2+ changes is unclear, but it has been suggested (61) that PAR-4-activated responses are not redundant, but reflect the necessity of temporally distinct and independent responses (see above). We also used intracellular Ca2+ measurements to determine the specificity of PAR interaction in epithelial cell function. In this study, we showed that thrombin and trypsin acted specifically on PAR-1 and PAR-2, respectively, and that frog PAR-1 and PAR-2 also acted specifically, consistent with data from previous studies. We also showed that human PAR-1 agonist peptide activated PAR-2 (32).

In conclusion, we have demonstrated that PAR agonist peptides induce IL-6, IL-8, and PGE2 release from human respiratory epithelium, and that these processes are accompanied by changes in intracellular calcium ion concentrations. Although the presence of all four PAR receptors in the lung epithelium was also confirmed, our study clearly demonstrated that the PAR-3 agonist peptide did not stimulate cytokine responses, consistent with functional data from other studies (21). The findings presented in this study also suggest that respiratory epithelium behaves similarly to other cell types expressing PAR. Our studies indicate that PARs are likely to be important in respiratory diseases, since a range of peptidases with the potential to activate PARs with concomitant release of proinflammatory mediators are known to be present in the lung of atopic individuals. However, whether this interaction results in inflammation and/or tissue repair is presently unclear. It is also possible that receptor expression up-regulation may occur, and, in this regard, we have recently shown that PAR-2 expression is up-regulated in bronchial biopsies from asthmatic subjects compared with control donors (63). Although the reason(s) for up-regulation is unclear, these observations, combined with data reported in this study, indicate that further examination of PAR in respiratory disease is warranted.


   Acknowledgments
 
We thank the Lotteries Commission, Western Australia, for the grant provided to Dr. Anthony Bakker for the purchase of the fluorescent microscope. We are grateful to Dr. L. F. Brass (University of Pennsylvania, Philadelphia, PA) for the generous gifts of the PAR-1 and PAR-2 Abs, and Sharon Redmond for photographic assistance.


   Footnotes
 
1 This work was supported by grants from the Asthma Foundation of Western Australia and the Australian National Health and Medical Research Council. Back

2 Address correspondence and reprint requests to Dr. Geoffrey A. Stewart, Department of Microbiology, University of Western Australia, 35 Stirling Highway, Crawley, Perth 6009, Western Australia, Australia. E-mail address: geoffrey{at}cyllene.uwa.edu.au Back

3 Abbreviations used in this paper: PAR, protease-activated receptor; HBEC, human bronchial epithelial cell; LHC, Lechner and La Veck medium; Indo, indomethacin; AM, acetoxymethylester. Back

Received for publication June 1, 2001. Accepted for publication January 9, 2002.


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 Materials and Methods
 Results
 Discussion
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