Human small airway epithelial cells (HSAEC) form the boundary between the external environmental allergens and the internal lung milieu. Mast cells are present in human lung tissue interspersed within the pulmonary epithelium and can secrete a host of pre- and newly formed mediators from their granules, which may propagate small airway inflammation. In this study, tryptase stimulation of HSAEC increased membrane-associated, calcium-independent phospholipase A2γ (iPLA2γ) activity, resulting in increased arachidonic acid and PGE2 release. These responses were inhibited by pretreating HSAEC with the iPLA2-selective inhibitor bromoenol lactone. The tryptase-stimulated PGE2 production was inhibited by treating HSAEC with the cyclooxygenase (COX)-1-selective inhibitor SC-560 and the nonselective COX inhibitor aspirin but not by the COX-2-selective inhibitor CAY10404, indicating that the early release of arachidonic acid is metabolized by constitutive COX-1 to form PGE2 in tryptase-stimulated HSAEC. Additionally, platelet-activating factor production and neutrophil adherence to tryptase-stimulated HSAEC was also increased. This complex response can set up a cascade of inflammatory mediator production in small airways. We speculate that selective inhibition of iPLA2γ-mediated phospholipid hydrolysis may prove beneficial in inflammatory airway diseases.
- prostaglandin E2
- arachidonic acid
- bromoenol lactone
the epithelium lining the respiratory tract in humans is composed of ciliated, nonciliated, and basal cells. These are the first cells to be exposed to an inhaled allergen (17). Damage to the epithelial cell lining results in cilial dysfunction, loss of cellular integrity, and penetration of allergens into the airways. Loss of epithelial integrity is a major feature of asthma pathogenesis and is thought to be an important contributor to the development of airway hyperresponsiveness (8). Resident mast cells in the pulmonary interstitium are important facilitators in the epithelial cell response to external injury. They secrete a host of mediators from the preformed granules including tryptase, chymase, and carboxypeptidase, which affect tissue remodeling and inflammatory cell recruitment (8, 25). In addition, mast cells also contain histamine, a vasoactive amine with potent effects on vascular permeability. Platshon and Kaliner (34) have concluded that histamine stimulation of human lung tissue leads to increased synthesis of PGF2α and cGMP in response to H-1 receptor stimulation and of cAMP through H-2 receptor stimulation. The rapid degranulation and release of tryptase following mast cell exposure to allergens provides a signal for activation of phospholipases, initiation of vascular change, and mobilization and recruitment of effector inflammatory cells (8, 17, 25, 34).
Previous studies from our laboratory (35, 36) have demonstrated that tryptase stimulation of endothelial cells can lead to activation of a calcium-independent phospholipase A2 (iPLA2), which hydrolyzes membrane phospholipids resulting in the stoichiometric production of a fatty acid, including arachidonic acid, and a lysophospholipid. In airway epithelial cells, arachidonic acid is further catabolized to eicosanoids, most typically PGE2. PGE2 acts on various E-prostanoid (EP) receptors and has been implicated in modulating human airway smooth muscle function. Activation of EP1 and EP3 receptors mediates airway smooth muscle contraction, whereas stimulation of EP2 receptors produces muscle relaxation (20).
The iPLA2 family of enzymes is made up of at least seven members (37). The first mammalian iPLA2 described was iPLA2β (group VIA PLA2; Ref. 14). In 2000, two independent laboratories identified a novel iPLA2, iPLA2γ (group VIB PLA2), which is constitutively membrane bound and possesses a COOH-terminal serine-lysine-leucine (SKL) peroxisomal targeting sequence (26, 28, 42). Multiple forms of iPLA2γ exist, and the expression of iPLA2γ is subject to rigorous control at many levels although the precise mechanisms for this are still not completely understood.
iPLA2γ protein expression has been confirmed in the mouse (28) and rat heart (27), rat liver (47), and human platelets (6) by immunoblot analysis. Overall, iPLA2γ expression appears to be widespread, but commercially available tools for the study of iPLA2γ (e.g., antibody, specific inhibitors, validated small interfering RNA, etc.) are only now becoming available. Both iPLA2β and iPLA2γ are sensitive to inhibition by bromoenol lactone (BEL) at low micromolar concentrations (26). However, other pharmacological PLA2 inhibitors, such as arachidonyl trifluoromethylketone (AACOCF3) and methyl arachidonyl fluorophosphonate (MAFP), which inhibit cytosolic iPLA2β at low micromolar concentrations (1, 21), do not inhibit microsomal iPLA2γ (10). Furthermore, Jenkins et al. (18) demonstrated that separation of racemic BEL into the R- and S-enantiomers provides a mechanism to discriminate between iPLA2γ and iPLA2β activity (i.e., iPLA2γ is ∼10-fold more sensitive to R-BEL than to S-BEL, and iPLA2β is 10-fold more sensitive to S-BEL than to R-BEL; Ref. 18).
This study examines the effect of tryptase stimulation on human small airway epithelial cell (HSAEC) PLA2 activity and the subsequent production of arachidonic acid, PGE2, and platelet activating-factor (PAF) and discusses the implications of these findings in small airway inflammation.
MATERIALS AND METHODS
HSAEC were obtained from Lonza Walkersville (Walkersville, MD). BEL, CAY10404, and SC-560 were obtained from Cayman Chemicals (Ann Arbor, MI). Recombinant human skin β-tryptase was obtained from Promega (Madison, WI). PX-18 was a gift from Richard Berney Associates (Bethesda, MD). CV3988 was purchased from Sigma-Aldrich (St. Louis, MO). All other reagents were purchased from Sigma-Aldrich.
Culture of epithelial cells.
HSAEC were grown to confluence in small airway basal medium (SABM; Lonza Walkersville) and incubated at 37°C with an atmosphere of 95% O2-5% CO2. Cells were passaged using subculture pack (Lonza Walkersville) in a 1:3 ratio. Cells from passages 3–4 were used for experiments.
Cells were grown to confluence in 100-mm culture dishes. At the end of each incubation period, media was removed and immediately replaced with ice-cold buffer containing, in mmol/l, 250 sucrose, 10 KCl, 10 imidazole, 5 EDTA, and 2 dithiothreitol, with 10% glycerol (pH 7.8). The cells were removed from the tissue culture plate by scraping, and the suspension was sonicated on ice for six bursts of 10 s. For separating the cells into membrane and cytosolic fractions, the sonicate was centrifuged at 14,000 g for 10 min. The supernatant was centrifuged at 100,000 g for 60 min to separate the membrane fraction (pellet) from the cytosolic fraction (supernatant). PLA2 activity was assessed by incubating enzyme (200 μg of cellular protein or 8 μg of membrane protein) with 100 μM (16:0, [3H]18:1) plasmenylcholine, (16:0, [3H]20:4) plasmenylcholine, (16:0, [3H]18:1) phosphatidylcholine, or (16:0, [3H]20:4) phosphatidylcholine (specific activity ∼150 dpm/pmol) in assay buffer containing 10 mM Tris, 10% glycerol, 4 mM EGTA, pH 7.0, at 37°C for 5 min in a total volume of 200 μl. For determining the sensitivity of membrane activity to R-BEL or S-BEL, the appropriate inhibitor was added to the isolated membrane fraction in the assay buffer for 10 min before the initiation of the activity assay. Reactions were initiated by adding the radiolabeled phospholipid substrate as a concentrated stock solution in ethanol. Reactions were terminated by the addition of 100 μl of butanol. The radiolabeled fatty acid released in the above reaction was isolated by application of 25 μl of the butanol phase to channeled Silica Gel G plates and then developed in petroleum ether-diethyl ether-acetic acid (70:30:1 vol/vol/vol). Results were quantified by liquid scintillation spectrometry.
Arachidonic acid release.
HSAEC were grown to confluence in 35-mm tissue culture dishes. Arachidonic acid release was determined by measuring [3H]arachidonic acid released into the surrounding medium from HSAEC prelabeled with 1 μCi of [3H]arachidonic acid (specific activity 100 Ci/mmol; PerkinElmer Life Sciences, Boston, MA) per culture dish for 18 h. Under these incubation conditions, 88% of the radiolabel was incorporated into the ethanolamine phospholipid pools of HSAEC. Analysis of the localization of incorporated exogenous deuterated arachidonic acid by mass spectrometry identified deuterated arachidonic acid in (16:0, 20:4), (18:0, 20:4), and (18:1, 20:4) plasmenylethanolamine, (16:0, 20:4) and (18:0, 20:4) phosphatidylethanolamine, and (16:0, 20:4) alkylacyl glycerophosphoethanolamine. Cells were washed three times with HEPES buffer containing, in mmol/l, 133.5 NaCl, 4.8 KCl, 1.2 CaCl2, 1.2 MgCl2, 1.2 KH2PO4, 10 HEPES (pH 7.4), 10 glucose, and 0.36% BSA and incubated at 37°C for 15 min before experimental conditions. At the end of the stimulation period, the surrounding medium was transferred to a scintillation vial, the remaining cells were lysed in 10% SDS, and the lysate was then transferred to a separate vial. Radioactivity in the medium and cells was quantified by liquid scintillation spectrometry. Arachidonic acid mobilized from cellular phospholipids was expressed as the percentage of total incorporated radioactivity.
HSAEC were grown to confluence in 16-mm tissue culture dishes. Cells were washed twice with HBSS containing, in mmol/l, 135 NaCl, 0.8 MgSO4, 10 HEPES (pH 7.6), 1.2 CaCl2, 5.4 KCl, 0.4 KH2PO4, 0.3 Na2HPO4, and 6.6 glucose. After washing, 0.5 ml of HBSS with 0.36% BSA was added to each culture well. HSAEC were then stimulated with the appropriate tryptase concentrations. The surrounding buffer was removed from the HSAEC after selected time intervals, and PGE2 release was measured immediately using an immunoassay kit (R&D Systems, Minneapolis, MN). The protein content for the HSAEC confluent monolayers was determined in three representative cell culture wells and was assumed to be constant between wells for each experiment.
HSAEC grown in 34-mm culture dishes (Corning) were washed twice with HBSS. Cells were incubated with 10 μCi of [3H]acetic acid per well for 20 min. After stimulation with tryptase, lipids were extracted from the cells using the method of Bligh and Dyer. The chloroform layer was concentrated by evaporation under nitrogen, resuspended in 9:1 CHCl3-CH3OH, applied to a silica gel 60 TLC plate, and developed in chloroform-methanol-acetic acid-water (50:25:8:4 vol/vol/vol/vol). The region corresponding to [3H]PAF was scraped, and radioactivity was quantified by liquid scintillation spectrometry. Loss of PAF during extraction and chromatography was corrected by adding a known amount of [14C]PAF as an internal standard.
HSAEC grown to confluence were incubated with BEL for 10 min and then removed from the tissue culture plate in 1.2 mM Ca2+ HEPES buffer and sonicated on ice. Cellular protein (25 μg) was incubated with 0.1 mM [acetyl-3H]PAF (10 mCi/mmol) for 30 min at 37°C. The reaction was stopped by the addition of acetic acid and sodium acetate. Released [3H]acetic acid was isolated by passing the reaction mixture through a C18 silica gel column (J. T. Baker, Phillipsburg, NJ), and eluted radioactivity was measured using a liquid scintillation counter.
Neutrophil isolation procedure.
Adult peripheral blood was collected (approved by Saint Louis University School of Medicine Institutional Review Board no. 12369) in vials containing 3.8% sodium citrate layered over Polymorphprep (Axis-Shield PoC AS, Oslo, Norway) and centrifuged at 500 g for 30 min. The top band at the sample/medium interface consisting of mononuclear cells and the lower band of polymorphonuclear cells were removed and washed with HBSS. Supernatant was discarded, and the cell pellet resuspended with 3 ml of 0.2% NaCl and incubated for 3 min at room temperature to lyse the red cells. Cells were again resuspended in HBSS and centrifuged at 175 g for 10 min at 4°C. Supernatant was removed, and cells resuspended in 5 ml of ice-cold HBSS. An aliquot was taken for a cell count using a hemacytometer.
Neutrophil adherence assay.
Neutrophils were resuspended in HBSS at 1 × 106 cells per milliliter. HSAEC were grown to confluence on a 12-well plate and washed with HBSS, and respective stimulants/inhibitors were added. CV3988 was added directly to the neutrophils when used. Five hundred microliters of neutrophils in suspension were added to each of the wells and incubated for 10 min at room temperature. Media and unbound neutrophils were removed and discarded. Plates were washed twice with Dulbecco's PBS. One milliliter of 0.2% Triton X-100 was added to each well to lyse adherent neutrophils and HSAEC. Cell lysates were scraped from the plate and transferred to an Eppendorf tube. A 500-μl aliquot of neutrophil suspension was added to 500 μl of 0.2% Triton X-100 and used as the theoretical maximal binding sample. Samples were sonicated (550 Sonic Dismembrator; Fisher Scientific, Pittsburgh, PA) for 10 s. To measure neutrophil peroxidase activity, 400 μl of cell lysate was transferred to a glass tube, 1 ml of PBS, 1.2 ml of HBSS + BSA, 200 μl of 3,3′-dimethoxybenzidine, and 200 μl of 0.05% H2O2 were added, and the mix was incubated for 15 min at room temperature. Two hundred microliters of 1% sodium azide (NaN3) was added to stop the reaction. The absorbance was then measured using a 4050 UV-Visible Spectrophotometer (Biochrom, Cambridge, England) at 450 nm.
Data were analyzed using the Student's t-test. P values of <0.05 were considered statistically significant; P values of <0.01 were considered highly statistically significant. ANOVA was used for comparison between multiple groups.
Effect of tryptase on PLA2 activity.
In initial experiments, we measured PLA2 activity in membrane and cytosolic subcellular fractions from HSAEC using (16:0, [3H]18:1) or (16:0, [3H]20:4) plasmenylcholine or phosphatidylcholine substrates in the presence (1 mM) or absence (4 mM EGTA) of calcium (Table 1). The majority of HSAEC PLA2 activity was measured in the absence of calcium (iPLA2), membrane-associated, and selective for arachidonylated (16:0, [3H]20:4) phospholipids (Table 1). Tryptase stimulation of HSAEC iPLA2 activity was significantly increased after 2 min and remained elevated for up to 30 min stimulation when activity was measured using (16:0, [3H]18:1) plasmenylcholine substrate in the absence of calcium (Fig. 1, open circles). However, no significant increase in iPLA2 activity was observed in tryptase-stimulated HSAEC when activity was measured using (16:0, [3H]18:1) phosphatidylcholine substrate (Fig. 1, closed squares). These data are consistent with our (35) previous finding in endothelial cells from other vascular beds and suggest that tryptase stimulates a plasmalogen-selective iPLA2. The tryptase-stimulated increase in iPLA2 activity was completely inhibited by pretreating the cells with the iPLA2-selective inhibitor BEL (2 μM, 10 min; Fig. 2, closed circles) but not by the secretory PLA2 (sPLA2) inhibitor PX-18 (2 μM, 10 min; Fig. 2, X). These data support our hypothesis that iPLA2 activity is increased in tryptase-stimulated HSAEC.
We further examined the specific iPLA2 isoform responsible for the increased activity measured in tryptase-stimulated HSAEC. We stimulated HSAEC with tryptase and then isolated the membrane and cytosolic subcellular fractions. iPLA2 activity was increased in the membrane fraction of tryptase-stimulated HSAEC, but cytosolic iPLA2 activity was not significantly altered (data not shown). Subsequently, the membrane fraction was incubated with increasing concentrations of R-BEL (selectively inhibits iPLA2γ) or S-BEL (selectively inhibits iPLA2β) for 10 min before the assay of iPLA2 activity. We found that iPLA2 activity was significantly inhibited by pretreatment with concentrations of R-BEL greater than 1 μM but was not inhibited by S-BEL until concentrations greater than 5 μM were used (Fig. 3). These data indicate that iPLA2γ is the major isoform present in HSAEC.
Effect of tryptase on arachidonic acid release from HSAEC.
Tryptase stimulation results in a fourfold increase in arachidonic acid release by 10 min compared with unstimulated cells (Fig. 4). Pretreatment with BEL (2 μM, 10 min) before tryptase stimulation completely inhibited tryptase-induced iPLA2 activation (Fig. 2) and arachidonic acid release (Fig. 4). Pretreatment of HSAEC with PX-18 (2 μM, 10 min) did not inhibit iPLA2 activity (Fig. 2) but inhibited tryptase-stimulated arachidonic acid release by ∼50% (Fig. 4). This indicates that arachidonic acid release in tryptase-stimulated HSAEC is dependent on activation of iPLA2 but that sPLA2 is also involved in tryptase-stimulated arachidonic acid release.
Effect of tryptase on PGE2 release in the presence of PLA2 inhibitors.
We stimulated HSAEC with tryptase and measured PGE2 release in the presence and absence of PLA2 inhibitors. Tryptase stimulation of HSAEC resulted in a significant increase in PGE2 release after 2 min (Fig. 5). Pretreatment with BEL (2 μM, 10 min) before tryptase stimulation completely inhibited the PGE2 release. Pretreatment with PX-18 (2 μM, 10 min) inhibited the tryptase-induced PGE2 release by ∼70% in these cells (Fig. 5).
Effect of tryptase on PGE2 release in the presence of cyclooxygenase inhibitors.
Arachidonic acid is metabolized by cyclooxygenase (COX)-1 and COX-2 to generate PGE2 in epithelial cells. Pretreatment with SC-560 (15 nM, 10 min, COX-1-selective inhibitor) or aspirin (1.5 nM, 10 min, COX-1 and COX-2 inhibitor) completely inhibited tryptase-stimulated PGE2 release from HSAEC (Fig. 6). However, pretreatment of cells with CAY10404 (1 nM, 10 min, a COX-2-selective inhibitor) had no effect. These data indicate that in the immediate phase of tryptase-stimulated PGE2 production it is the constitutive COX-1 isoform that is responsible for metabolizing iPLA2-catalyzed arachidonic acid release to generate PGE2.
Effect of tryptase on PAF production.
The lysophospholipid generated as a consequence of removal of sn-2 fatty acid from the phospholipid by iPLA2 is rapidly acetylated to PAF. PAF is a potent proinflammatory lipid metabolite that can result in disruption of the airway epithelial defense barrier. Tryptase stimulation of HSAEC caused an increase in PAF production that was observed until 30 min. This effect was attenuated by pretreating the cells with BEL (Fig. 7) indicating that tryptase-induced PAF production in HSAEC is at least in part due to iPLA2 activation. Treatment of HSAEC with concentrations of BEL up to 10 μM did not inhibit PAF-acetylhydrolase activity (5.72 nmol·mg protein−1·min−1 in untreated cells vs. 6.03 nmol·mg protein−1·min−1 in cells treated with 10 μM BEL for 10 min, n = 3 separate cell cultures).
Effect of tryptase on neutrophil adherence.
Previous studies from our laboratory have demonstrated significant increases in neutrophil adherence to tryptase-stimulated coronary artery endothelial cells. We examined whether tryptase stimulation would have a similar effect on the adherence of neutrophils to the HSAEC monolayer. Tryptase stimulation (20 ng/ml, 10 min) led to a significant increase in neutrophil adherence (Fig. 8). We also saw that pretreatment of neutrophils with a PAF receptor antagonist, CV3988, reduced the percentage of adherent neutrophils by 80% (from 14.77% ± 0.27% to 3.25% ± 0.25%, n = 4) demonstrating the importance of PAF-PAF receptor interactions in the adherence of these cells.
Activation of mast cells can release several mediators from their granules that are responsible for a variety of inflammatory allergic disorders such as allergic rhinitis, asthma, dermatitis, and anaphylactic responses. In the lung, mast cells are found widely distributed in pleura, peribronchial regions, alveolar septa adjacent to nerves and blood vessels, upper and lower respiratory epithelium, and within the bronchial lumen (8). Tryptase released from mast cell granules has widespread effects on cellular proteins that can contribute to the pathogenesis of pulmonary inflammation in large and small airways (41). Tryptase degrades vasoactive intestinal polypeptide, an endogenous bronchodilator, generates bronchoconstrictor kinins, and activates prekallikrein (9). It also increases airway smooth muscle responsiveness to histamine in vitro and airway hyper responsiveness in vivo.
Airway inflammation in asthma is widely considered to be due to changes in the large airways. However, there is a significant patient population that does not respond to currently available antiasthmatic therapeutic regimens. In such cases, it is believed that the small airway (<2 mm in diameter) could be responsible for the residual inflammation. However, there are technical difficulties associated with evaluation of distal lung tissue. Only a small number of studies, using tissue obtained at the time of tumor resection or alveolar tissue from transbronchial biopsies, have evaluated distal lung inflammatory cell distribution and function (31, 38, 40).
Small airways consist of the respiratory bronchiole and terminal bronchioles that are devoid of cartilage and mucous secreting glands (12). Pathological changes in the small airways may proceed long before clinical symptoms of pulmonary inflammation become apparent. A hallmark of airway inflammation is widespread damage to the epithelium. There is infiltration by neutrophils and activated eosinophils (23). Epithelial sloughing occurs during exacerbation of bronchial asthma and is due to a possible defect in adherence to the basement membrane (5). The degree of airway hyperresponsiveness has been postulated to correlate with the loss of airway epithelial cells (45).
The airway epithelium can serve as both a target and effector cell in propagating the inflammatory response. On encountering allergens, the airway epithelium reacts by increasing mucus secretion, ciliary beat frequency, and changes in the ion transport/barrier function. The inflammatory metabolites released by the epithelium can act in an autocrine or paracrine fashion to exert widespread effects. Previous studies have demonstrated that airway epithelium can release IFN-γ and TNF-α. These in turn can lead to production of several secondary mediators such as prostanoids (PGE2), nonprostanoid mediators including epithelium derived relaxant/inhibitory factor, reactive oxygen species, and nitric oxide. PGE2 can have both pro- and anti-inflammatory effects. Inhaled PGE2 abolishes exercise-induced bronchoconstriction (29). It also inhibits bronchial hyperresponsiveness to inhaled allergen and attenuates antigen-induced eosinophilic inflammation (11, 33). Since PGE2 is a potent vasodilator, it potentiates the edema induced by other mediators (46). In addition, it reinforces the T helper type 2 (TH2) responses by increasing T cell polarization and release of IL-10 while inhibiting the release of IL-12 (a TH1-polarizing cytokine) (13).
To the best of our knowledge, this is the first study that demonstrates that mast cell tryptase can activate membrane-associated iPLA2γ in small airway epithelial cells resulting in arachidonic acid release, which is metabolized by COX-1 to generate PGE2. Pretreating the cells with BEL led to complete inhibition of these responses, whereas PX-18 led to partial inhibition of arachidonic acid release and PGE2 generation. This suggests that iPLA2 is required for the production of arachidonic acid in the immediate response to tryptase in airway epithelial cells but that sPLA2 may also be involved to a lesser degree.
There are three main classes of PLA2 that coexist in mammalian cells, secretory, cytosolic, and calcium-independent PLA2. They are activated in a temporal sequence and exert their effect over varying lengths of time depending on the stimulus and cell type (30). The cyclooxygenases convert arachidonic acid to an intermediate precursor PGH2. The release of arachidonic acid and formation of PGH2 is the crucial rate-limiting step for prostaglandin synthesis. In the early phase of prostaglandin biosynthetic response, increased arachidonic acid production by iPLA2 activation is followed by oxidation by constitutive COX-1 and prostaglandin biosynthesis (32).
Several published studies have described a possible role for iPLA2 in the generation of eicosanoids. IgG receptors (FcεR) have been demonstrated to functionally couple to iPLA2β for the release of arachidonic acid and the production of leukotriene B4 and PGE2 (43). Murakami et al. (32) transfected various PLA2 and COX enzymes into human embryonic kidney cells and observed that ionophore-induced immediate PGE2 generation was linked to iPLA2β and COX-1 activity. They suggested that iPLA2 releases arachidonic acid in closer proximity to COX-1 than COX-2 and that iPLA2-derived arachidonic acid is somehow inaccessible to COX-2 (35). We report that immediate release of PGE2 from tryptase-stimulated HSAEC is dependant on activation of membrane-associated iPLA2γ and subsequent metabolism of arachidonic acid by COX-1, further supporting this hypothesis.
Although intracellular PLA2 isoforms may be directly coupled to COX in the cell for eicosanoid production, several studies suggest that immediate eicosanoid production involves cytosolic PLA2 (cPLA2) and sPLA2 with cPLA2 being the activator of the response but sPLA2 providing the bulk of arachidonic acid (22). Since tryptase-stimulated PGE2 production in HSAEC was significantly inhibited by pretreatment with BEL and partly by PX-18, it is possible that a similar interaction between iPLA2 and sPLA2 is involved in arachidonic acid release. Cross talk between cPLA2 and sPLA2 has been previously demonstrated in P388D1 macrophages in which intracellular arachidonic acid release by cPLA2 regulates the accessibility of sPLA2 to its substrate in the membrane (3). A similar mechanism, at least in part, may exist between iPLA2 and sPLA2 in HSAEC since PX-18 inhibits tryptase-stimulated arachidonic acid release and PGE2 production but does not inhibit iPLA2 activity.
In our studies, we also saw increased adherence of neutrophils to tryptase-stimulated HSAEC mediated by PAF. Holtzman et al. (16) have previously demonstrated PAF production by airway epithelial cells. PAF activates leukocytes and platelets via specific cell surface receptors, induces leukocyte chemotaxis, and stimulates preferential migration of eosinophils into the airway (2). It can also induce airway smooth muscle contraction and hyperreactivity in healthy subjects (19). Airway epithelial cells express several leukocyte adhesion molecules, and the receptors for these ligands are regulated by PAF (24, 44).
Infiltration and adherence of neutrophils occurs during asthma and chronic bronchitis and is also a hallmark of cystic fibrosis (39). Tryptase is known to have mitogenic effect on airway smooth muscles. In conjunction with increased neutrophil adherence, it may set up a cycle of inflammation that can induce airway remodeling and compromise lung function (7). In light of the close proximity of HSAEC and the interstitial mast cells, the effects of tryptase seen in the above-mentioned studies can be an important pathway in propagating small airway inflammation.
The epithelium is usually regarded as a target for inflammatory mediators, but this study demonstrates that it is capable of initiating and sustaining an inflammatory response. Once targeted, the epithelium sets a cascade of inflammatory events, which can be responsible for activation of inflammatory cells, stimulation of smooth muscle, and fibroblast proliferation. Molecular mechanisms that regulate COX and lipoxygenase production in airway epithelium provide an important therapeutic target. In asthma, PLA2 activation triggered by mast cell tryptase may contribute to the propagation of inflammation via the production of several membrane phospholipid-derived metabolites. Thus the development of a specific PLA2 inhibitor has the potential to be a valuable therapeutic tool for treating early airway inflammation that contributes further to the pathogenesis of asthma.
This work was supported in part by National Heart, Lung, and Blood Institute Grant HL-68588 (J. McHowat) and the American Heart Association Heartland Affiliate 0610118Z (P. Rastogi).
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