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J Appl Physiol 91: 995-1003, 2001;
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Vol. 91, Issue 2, 995-1003, August 2001

HIGHLIGHTED TOPICS
Signal Transduction in Smooth Muscle
Selected Contribution: Tryptase-induced PAR-2-mediated Ca2+ signaling in human airway smooth muscle cells

Patrick Berger, J. Manuel Tunon-De-Lara, Jean-Pierre Savineau, and Roger Marthan

Laboratoire de Physiologie Cellulaire Respiratoire, Inserm E9937, Université Bordeaux 2, 33076 Bordeaux, France


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Tryptase, the major mast cell product, is considered to play an important role in airway inflammation and hyperresponsiveness. Tryptase produces different, sometimes opposite, effects on airway responsiveness (bronchoprotection and/or airway contraction). This study was designed to examine the effect of human lung tryptase and activation of protease-activated receptor (PAR)-2 by synthetic activated peptide (AP) SLIGKV-NH2 on Ca2+ signaling in human airway smooth muscle (HASM) cells. Immunocytochemistry revealed that PAR-2 was expressed by HASM cells. Tryptase (7.5-30 mU/ml) induced a concentration-dependent transient relative rise in cytoplasmic Ca2+ concentration ([Ca2+]i) that reached 207 ± 32 nM (n = 10) measured by indo 1 spectrofluorometry. The protease inhibitors leupeptin or benzamidine (100 µM) abolished tryptase-induced [Ca2+]i increase. Activation of PAR-2 by AP (1-100 µM) also induced a concentration-dependent transient rise in [Ca2+]i, whereas the reverse peptide produced no effect. There was a homologous desensitization of the [Ca2+]i response on repeated stimulation with tryptase or AP. U-73122, a specific phospholipase C (PLC) antagonist, xestospongin, an inositol trisphosphate (IP3)-receptor antagonist, or thapsigargin, a sarcoplamic Ca2+-ATPase inhibitor, abolished tryptase-induced [Ca2+]i response, whereas Ca2+ removal, in the additional presence of EGTA, had no effect. Calphostin C, a protein kinase C inhibitor, increased PAR-2 [Ca2+]i response. Our results indicate that tryptase activates a [Ca2+]i response, which appears as PAR-2 mediated in HASM cells. Signal transduction implicates the intracellular Ca2+ store via PLC activation and thus via the IP3 pathway. This study provides evidence that tryptase, which is increasingly recognized as an important mediator in airway inflammation and hyperresponsiveness, is also a potent direct agonist at the site of airway smooth muscle.

calcium release; protease; protease-activated receptors; cytoplasmic calcium concentration cell culture


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

PROTEASES HAVE MULTIPLE BIOLOGICAL roles, and certain proteases possess biological activity that is receptor mediated via protease-activated receptors (PARs), an expanding family of G-protein-coupled receptors (43). Although PARs are only a small component of this large family, they are functionally important because they regulate inflammation, responses to injury, growth, and development (12). Activation of PARs is different from that of other seven-transmembrane-domain G-protein-coupled receptors. Proteases cleave PARs within the extracellular NH2-terminal domain, exposing a new NH2 terminus that acts as a tethered ligand by binding to extracellular domains of the receptor and thereby activating the cleaved receptor molecule (43). Synthetic peptides corresponding to the tethered ligands of PARs activate the corresponding PARs (16).

Most of the biological effects of tryptase, a major secretory granule protease of human mast cells, are also receptor mediated via the PAR subtype, PAR-2 (24). It is noteworthy that, on the basis of trypsin experiments, tryptase (a trypsinlike protease) may also activate PAR-4 (45). Tryptase is considered to play an important role in airway inflammation (40) and hyperresponsiveness (7). Moreover, since the pioneering work of Sekizawa et al. (37) in dog isolated airways, it has been demonstrated that tryptase also potentiates contraction of human isolated airways in spontaneously sensitized (18) or even nonsensitized lungs (2).

PAR-2 has been localized to both animal (30) and human airways (11). Activation of PAR-2 produces several different, sometimes opposite, effects on airway responsiveness. On the one hand, activation of PAR-2 can initiate powerful bronchoprotection in the airways (9). This effect appears mediated by activation of epithelial PAR-2 and resembles that of the endothelium-dependent relaxation of blood vessels (16). On the other hand, activation of PAR-2 can produce airway contraction. In guinea pig airways, trypsin induced bronchoconstriction in vivo, which appears to depend on both direct smooth muscle contraction and indirect mechanisms (30), one being a release of sensory neurokinins, as demonstrated in response to trypsin stimulation (6).

A direct PAR-2-mediated contractile effect of mast cell tryptase has been identified in rat colonic smooth muscle cells (10). This effect was ascribed to a tryptase-induced increase in the cytoplasmic free Ca2+ concentration ([Ca2+]i) (10). Regarding airways, alpha -thrombin, another protease that preferentially activates PAR-1, has been shown to both contract human airway smooth muscle (HASM) (15) and increase [Ca2+]i (27). However, although implicated in airway inflammation and hyperresponsiveness (7), to the best of our knowledge, tryptase-induced PAR-2-mediated signal transduction in HASM has not been examined so far.

The present study was thus designed to examine tryptase-induced Ca2+ signaling in isolated HASM cells. For this purpose, we have examined the effects of both human mast cell tryptase and synthetic peptides corresponding to the tethered ligand domain, i.e., the agonist peptide (AP) SLIGKV-NH2 and the reverse peptide (RP) VKGILS-NH2, using indo 1 microspectrofluorometry in both freshly isolated and cultured single HASM cells.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Purification and characterization of tryptase. Human mast cell tryptase was purified from lung tissue obtained post mortem. The tissue (400-500 g) was chopped finely and homogenized, after which it was first incubated in a low-salt buffer and then subjected to a high-salt extraction procedure, as described previously (44). The supernatant was filtered through a microfiber membrane, dialyzed against distilled water (24 h, 4°C), and subjected to heparin-agarose affinity chromatography, equilibrating with a low-salt buffer. Fractions were eluted using a NaCl gradient between 0.4 and 1.5 M in 10 mM MES (Sigma Chemical, St. Quentin Fallavier, France) buffer. Tryptase-rich fractions were then subjected to a benzamidine-agarose affinity chromatography, equilibrating with a high-salt buffer (2 M NaCl, 10 mM MES). Fractions were eluted using 0.15 M benzamidine and concentrated using an Amicon concentrator with a YM30 membrane that separated tryptase from benzamidine. Filtrates were then applied to a Sephacryl S-300 gel filtration column equilibrated with high-salt buffer (2 M NaCl, 10 mM MES), and tryptase-rich fractions were again concentrated, passed through a 0.22-µm membrane filter, and stored at -80°C. The purity of the tryptase samples was confirmed by SDS-PAGE with 10% reducing gels. The preparation employed in these studies appeared as a single band on silver staining with a molecular mass of ~34 kDa. The identity of the purified protein was confirmed by immunoblotting with the tryptase-specific monoclonal antibody AA5 (44).

Enzyme assay. Tryptase activity was determined using the synthetic peptide substrate N-benzoyl-DL-Arg-p-nitroanilide (BAPNA), adding 10 µl of enzyme to 90 µl of 20 mM Tris buffer containing 1 M glycerol and 7.77 mM BAPNA. Absorbance was measured at 410 nm in an ELISA plate reader. Protein concentration was measured spectrophotometrically at 280 nm, using the coefficient of extinction of Smith et al. (39). The specific activity of the tryptase preparation employed was 2.9 U/mg, where 1 unit represents the amount of tryptase required to hydrolyze 1 µmol of substrate per minute at 25°C. The enzyme preparation was found to be 100% active by titration with the substrate 4-methylumbelliferyl p-guanidinobenzoate (Sigma Chemical). No chymase or elastase activity was found in these samples, using the chromogenic substrates N-succinyl-L-Ala-L-Ala-L-Pro-L-Phe-p-nitoanilide and N-succinyl-L-Ala-L-Ala-L-Pro-L-Val-p-nitoanilide (both from Sigma Chemical), respectively.

Tissue preparation. Tissues were collected after lung resection for bronchial carcinoma (n = 23) and immediately transferred to the laboratory in sterile DMEM (GIBCO BRL Life Technologies, Cergy Pontoise, France). As in previous studies, specimens were selected from patients whose lung function was within a normal range, i.e., whose forced expiratory volume in 1 s and total lung capacity were above 80% of predicted (2, 22). From a macroscopically tumor-free part of the specimen, segments of human bronchus (3rd to 4th division) were carefully dissected under a dissecting microscope. After removal of adhering fat, parenchyma, epithelium, and submucosal tissue, the smooth muscle bands were cut into squares measuring 1-2 mm2 to be either enzymatically dissociated or cultured.

Cell dissociation. Freshly isolated HASM cells were enzymatically dissociated as previously described (22). Briefly, the smooth muscle squares were incubated for 10 min in zero-Ca2+ physiological saline solution (PSS, composition given below) and then incubated in zero-Ca2+ PSS containing 500 µg/ml collagenase type I (Worthington Biochemical, Freehold, NJ), 350 µg/ml pronase (Sigma Chemical), 31.25 µg/ml elastase type III (Sigma Chemical), and 1 mg/ml soybean trypsin inhibitor (Sigma Chemical) at 4°C for 14 h. After this time, the solution was removed, and the bronchial pieces were incubated again in a fresh enzyme-free solution and triturated with a fire-polished Pasteur pipette to release cells. Cells were stored on glass coverslips at 4°C in PSS containing 0.8 mM Ca2+ and used on the same day.

Cell culture. Smooth muscle explants were cultured in six-well culture plates in a humidified atmosphere at 37°C with 5% CO2. HASM cells were maintained in DMEM containing 10% (vol/vol) FCS (GIBCO), supplemented with 2 mM L-glutamine (GIBCO), 1 mM sodium pyruvate (Sigma Chemical), 1% (vol/vol) nonessential amino acid mixture (Sigma Chemical), 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B (antimycotic-antibiotic solution, GIBCO). The medium was changed every 48-72 h. After 6-8 wk, confluent cells were rinsed twice with HBSS (GIBCO) and then passaged with trypsin-EDTA (GIBCO). Cells were seeded on glass coverslips at a density of 105 cells/ml. Only cells at passages 2-4 were used for this study. After 14 h, growth was arrested by incubating the cells for 24-48 h with serum-free DMEM supplemented with 10 µg/ml insulin, 5.5 µg/ml transferrin, 5 ng/ml selenium, 0.5 µg/ml BSA, 4.7 µg/ml linoleic and oleic acid (ITS solution, Sigma Chemical), 2 mM L-glutamine (GIBCO), 1 mM sodium pyruvate (Sigma Chemical), 1% (vol/vol) nonessential amino acid mixture (Sigma Chemical), 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B (antimycotic-antibiotic solution, GIBCO).

Immunocytochemistry. To assess the purity of the cells, an immunocytochemical method was employed using an indirect immunofluorescence technique. Cells of varying passage number were growth arrested as described above. After 24 h, cells were rinsed twice in PBS (GIBCO) and fixed with cold methanol for 20 min. Nonspecific staining was blocked using PBS containing 3% BSA (GIBCO) for 30 min. Monoclonal antibodies diluted in PBS with 1% BSA, including anti-alpha -smooth muscle actin (1:200, Sigma Chemical), anti-smooth muscle myosin (1:200, Sigma Chemical), anti-cytokeratin 18 (1:500, Sigma Chemical), anti-factor VIII (1:25, Dako), anti-fibroblast surface protein (1:100, Sigma Chemical), or anti-human PAR-2 (1:100, Santa Cruz Biotech, Santa Cruz, CA), were incubated for 1 h. Control slides were treated similarly, omitting the primary monoclonal antibodies or using an unrelated antibody (mouse or goat immunoglobulin G, Sigma Chemical). After cells were rinsed with PBS containing 0.05% Tween 20 (Sigma Chemical), the cells were incubated for 1 h with FITC-conjugated anti-mouse immunoglobulins (Dako), diluted 1:20 except for anti-PAR-2 for which the secondary antibody was FITC-conjugated anti-goat immunoglobulin (Sigma Chemical) diluted 1:400. Counterstaining was performed using 2 µg/ml propidium iodide (Sigma Chemical). Slides were mounted with a drop of 10% glycerol in PBS and observed under a Diastar fluorescence microscope.

Fluorescence measurement and estimation of [Ca2+]i. Changes in [Ca2+]i were monitored fluorometrically by use of the Ca2+-sensitive probe indo 1 as described previously (32). Briefly, cells were loaded with indo 1 (Calbiochem, Meudon, France) by incubation in PSS containing 1.25 µM indo 1 penta-acetoxymethyl ester (indo 1-AM) for 25 min at room temperature and then washed in PSS for 25 min. The coverslip with attached cells was then mounted in a perfusion chamber and continuously perfused. The recording system included a Nikon Diaphot inverted microscope fitted with epifluorescence (Nikon France, Charenton-le-pont, France). A single cell among those on the coverslip was tested through a window slightly larger than the cell, created by a pinhole placed in the light beam. The cell was illuminated at 360 nm, and emitted light was counted simultaneously at 405 nm and 480 nm by two photomultipliers (P100, Nikon). Voltage signals at each wavelength were stored in an IBM-PC computer for subsequent analysis. The fluorescence ratio (405 nm/480 nm) was calculated on-line and displayed with the two voltage signals on a monitor. [Ca2+]i was estimated from the 405-to-480-nm ratio (14) using a calibration for indo 1 determined within cells (32).

Solutions and ejection of agonists. The normal PSS contained (in mM) 130 NaCl, 5.6 KCl, 1 MgCl2, 2 CaCl2, 11 glucose, and 10 HEPES, pH 7.4 with NaOH. Ca2+-free PSS was prepared by replacing CaCl2 with 0.4 mM EGTA.

Agonists were then applied to the recorded cell by pressure ejection from a glass pipette located close to the cell for the period indicated on the records. Purified human lung tryptase was added in the presence of heparin (in a weight ratio of 1:1) to stabilize enzymatic activity (35). To investigate dependency on an intact catalytic site, tryptase (with added heparin) was incubated in the presence or absence of the enzymatic inhibitors leupeptin (100 µM, Sigma Chemical) and benzamidine (100 µM, Sigma Chemical). The effect of heat treatment was also investigated, by heating tryptase at 56°C for 60 min. Some experiments were also performed using bovine trypsin. The potential involvement of PAR-2 in HASM cell Ca2+ response was investigated using the AP SLIGKV-NH2 and, as a control, the RP VKGILS-NH2 (both synthesized by MWG-Biotech). Bovine trypsin (Sigma Chemical), a serine protease that cleaves PAR-2, was also tested. Conventional airway smooth muscle agonists such as histamine (Sigma Chemical), ACh (Sigma Chemical), and caffeine (Merck, Darmstadt, Germany) acted as positive controls. In control experiments, no change was observed in [Ca2+]i during the ejections of PSS or heparin alone.

The effect of incubating cells with various inhibitors of signal transduction pathways was also studied. Pertussis toxin (50 ng/ml, Sigma Chemical) was added for 18-24 h before tryptase challenge. Neomycin (1 µM), thapsigargin (TG, 1 µM), U-73122, and U-73343 (5 µM) (all from Sigma Chemical) were incubated for 1 h before challenge. Calphostin C (0.5 µM, Sigma Chemical) and xestospongin C (10 µM, Calbiochem) were incubated 20 min before challenge.

Generally, each record of [Ca2+]i response to each agonist alone or in the presence of an additional substance was obtained from a different cell. In another set of experiments, the same cell was challenged several times with one or several agonists. Each stimulation was separated from the next by a sufficient time lag to allow for the refilling of intracellular Ca2+ stores (generally 5 min). Each type of experiment was repeated for the number of patient-derived smooth muscle cell lines indicated in the text. Experiments were done at room temperature (22-25°C).

Statistical analysis. Results are expressed as means ± SE, with n referring to the number of different lung specimens used to obtain the cells. Each experimental condition was tested in at least 10 different cells prepared from each lung specimen. Significance was tested by one-way ANOVA at a P value of <0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell purity and immunodetection of PAR-2. All HASM cells stained positively for smooth muscle actin and myosin (Fig. 1, A and B). There was no apparent variation in staining intensity between cells of different passage numbers. HASM cells also expressed PAR-2 (Fig. 1, C and D). No immunostaining was seen with antibodies specific for cytokeratin (Fig. 1E), factor VIII, or fibroblast (data not shown).


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  		Fig. 1.   Immunocytochemistry performed in human cultured airway smooth muscle with monoclonal antibodies directed against smooth muscle actin (A), myosin (B), protease-activated receptor (PAR)-2 (C and D), cytokeratin (E), or negative control (goat IgG, F). Cells were specifically stained in green with FITC-conjugated secondary antibody.

[Ca2+]i response to ACh, histamine, and trypsin. In both freshly isolated and cultured HASM cells, ACh, histamine, or trypsin induced a [Ca2+]i rise. There was no significant difference in terms of [Ca2+]i resting value and relative [Ca2+]i peak in response to the three compounds between both cell types (Table 1). However, the percentage of responding cells to trypsin was higher in quiescent cultured cells than in freshly isolated HASM cells (Table 1). This difference was even higher for lower concentrations of trypsin: 90 and 49% for freshly isolated and cultured HASM cells, respectively, for 10-8 M trypsin, and 88 and 35% for 10-9 M trypsin. For this reason, subsequent experiments were conducted in cultured HASM cells.

                              
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  		Table 1.   Effect of acetylcholine, histamine, and trypsin on intracellular Ca2+

Characteristics of the tryptase-induced [Ca2+]i response in quiescent cultured HASM cells. Single HASM cells responded to application of exogenous human tryptase (7.5-30 mU/ml, 30 s) by a transient rise in [Ca2+]i, the amplitude of which was concentration dependent (Fig. 2, A-C). The mean maximal amplitude of the [Ca2+]i peak was 297 ± 31 nM (n = 12) above a mean resting value of 91 ± 4 nM. The mean time between the beginning of the ejection and the peak was 26.4 ± 5.1 s, whereas it was shorter for the conventional agonists, ACh and histamine (14.8 ± 1.6 and 9.6 ± 1.3 s, P < 0.05, respectively).


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  		Fig. 2.   Effect of human purified tryptase on cytoplasmic Ca2+ concentration ([Ca2+]i) in human cultured airway smooth muscle cells. Typical [Ca2+]i responses evoked by a short (30 s) stimulation with 7.5 (A), 15 (B), or 30 mU/ml of tryptase in the absence (C) or presence (D) of 100 µM leupeptin. Each trace is representative of experiments performed in 40-50 cells.

The protease inhibitors leupeptin or benzamidine (100 µM), which reduced the enzymatic activity of tryptase to 98 and 50%, respectively, also abolished tryptase-induced [Ca2+]i increase even when tryptase was ejected for a longer period of time (Fig. 2D). We did verify that leupeptin, the most potent tryptase inhibitor, altered the Ca2+ response of HASM cells to neither ACh nor histamine (data not shown). In this connection, heat inhibition of tryptase activity (1 h at 56°C) also abolished tryptase-induced HASM cell Ca2+ response (data not shown).

Involvement of PAR-2 in tryptase-induced [Ca2+]i response. The activation of PAR-2 by the AP (SLIGKV, 1-100 µM) induced a concentration-dependent transient rise in [Ca2+]i (Fig. 3, A and B), whereas the RP (VKGILS), which does not activate PAR-2, produced no effect up to 100 µM (Fig. 3C). The mean maximal AP-induced [Ca2+]i peak reached 331 ± 23 nM (n = 8) from a mean resting [Ca2+]i value of 106 ± 6 nM, and the mean time between the beginning of the ejection and the peak was 23.3 ± 2.3 s. These results are close to those obtained with tryptase.


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  		Fig. 3.   Effect of PAR-2 activation on the [Ca2+]i in cultured human airway smooth muscle cells. Typical [Ca2+]i responses evoked by a short (30 s) stimulation with 1 (A) or 100 µM (B) agonist peptide (AP; SLIGKV), 100 µM reverse peptide (RP; VKGILS) (C), or 100 nM trypsin (D). Each trace is representative of experiments performed in 80-100 cells.

PAR-2 can also be activated by trypsin. In HASM cells, trypsin (1-100 nM) induced a concentration-dependent transient rise in [Ca2+]i (Fig. 3D). Again, the characteristics of the trypsin-induced [Ca2+]i response were close to those of tryptase. The amplitude of the maximal [Ca2+]i peak was 312 ± 23 nM, from a resting value of 103 ± 8 nM (n = 8), and the mean time between the beginning of the ejection and the peak was 25.3 ± 1.6 s. As for tryptase, the trypsin-induced Ca2+ response was reduced by preincubation with the protease inhibitor benzamidine (not shown).

A characteristic feature of PAR is the occurrence of homologous desensitization related to the mechanism of activation of these receptors. When a single HASM cell was stimulated twice with tryptase, the amplitude of the Ca2+ response was significantly decreased (345 ± 27 vs. 101 ± 35 nM, n = 4; Fig. 4A). A similar homologous desensitization of Ca2+ responses was observed when the subsequent stimulation was performed with trypsin (331 ± 38 vs. 112 ± 24 nM, n = 4; Fig. 4B) or the AP SLIGKV (174 ± 59 vs. 74 ± 14 nM, n = 3; not shown). Two subsequent stimulations by the AP SLIGKV also desensitized the PAR-2-mediated [Ca2+]i response (381 ± 62 vs. 180 ± 30 nM, n = 7, Fig. 4C). In all of the above-described experiments, specificity of PAR-2-mediated desensitization of [Ca2+]i response was assessed by the fact that a third subsequent stimulation of the cells with a conventional agonist (ACh or histamine) produced a Ca2+ increase similar to that for prior PAR-2 activation (not shown).


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  		Fig. 4.   Effect of desensitization of PAR-2 with human purified tryptase (A and B) or AP (SLIGKV) (C) on [Ca2+]i in cultured human airway smooth muscle cells. Cells were stimulated twice with 30 mU/ml of tryptase (A), 30 mU/ml of tryptase and then with 100 nM of trypsin (B), or twice with 100 µM AP (C). Each trace is representative of experiments performed in 30-50 cells.

Signal transduction in PAR-2-mediated [Ca2+]i response. A variety of pharmacological tools were used to identify the transduction pathway linking PAR-2 activation to the [Ca2+]i response. Preincubation of cells with pertussis toxin (50 ng/ml for 24 h) did not alter the tryptase response (Table 2). In contrast, U-73122 (5 µM for 1 h), a specific phospholipase C (PLC) antagonist, abolished the tryptase-induced [Ca2+]i response (Table 2). U-73122 also abolished the [Ca2+]i response produced by the alternative PAR-2 agonists, trypsin and the AP SLIGKV (Table 2). [Ca2+]i responses to conventional agonists, ACh and histamine, were also inhibited by U-73122. Similar results were obtained with neomycin, another PLC antagonist (not shown). The negative control U-73343, which does not inactivate PLC, did not significantly alter either the tryptase-induced or other PAR-2 agonist-induced responses, nor did it significantly alter responses to ACh or histamine (Table 2). To evaluate the role of the metabolites produced by PLC activation, xestospongin C [an inositol trisphosphate (IP3)-receptor antagonist] and calphostin C [a protein kinase C (PKC) inhibitor] were preincubated for 20 min. Xestospongin C inhibited the [Ca2+]i response to tryptase, trypsin, AP, and conventional agonists (ACh and histamine) (Table 2). Calphostin C did not inhibit any of these responses but surprisingly increased tryptase, trypsin, and AP-induced [Ca2+]i responses (Table 2); however, it did not modify ACh or histamine responses.

                              
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  		Table 2.   Signal transduction mechanisms involved in the cultured HASM cell Ca2+ response

The Ca2+ source implicated in the PAR-2-mediated rise store has been investigated by means of removal of external Ca2+ and by using TG, a sarcoplasmic Ca2+-ATPase inhibitor (13) at a concentration of 1 µM for 30 min. TG completely inhibited tryptase-induced Ca2+ responses, whereas Ca2+ removal, in the additional presence of EGTA, did not alter the tryptase-induced [Ca2+]i rise (Table 2). Again, similar results were obtained with alternative PAR-2 activators as well as with ACh or histamine (Table 2).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The present study indicates that human mast cell tryptase has a direct effect on isolated HASM cells. Tryptase activates a [Ca2+]i response that appears to be PAR-2 mediated. In accordance with the mechanism of activation of PAR, the tryptase-induced [Ca2+]i rise is delayed compared with more conventional agonists such as ACh or histamine, which also act at G-protein-coupled receptors. Nevertheless, signal transduction mechanisms downstream of the receptor level are similar for tryptase and ACh or histamine. PAR-2 activation mobilizes the intracellular Ca2+ store (presumably the sarcoplasmic reticulum) via phosphoinositide PLC activation and thus via the IP3 pathway.

Our data provide some evidence for a direct action of tryptase on HASM, because results were obtained on pure, isolated cells, as confirmed by immunocytochemistry. Most of the experiments were conducted on cultured cells once it was verified that these cells exhibited agonist-induced Ca2+ responses similar to those of freshly isolated cells (Table 1). In performing this comparison, the main difference that we observed was a decrease in percentage of cells responding to PAR agonist in freshly isolated cells. This observation is not unexpected because PARs are likely to be even more sensitive to the enzymatic dissociation than any other membrane receptor (42).

The tryptase-induced [Ca2+]i response appears to be PAR-2 mediated for the following reasons. First, irrespective of the PAR subtype, it should be noted that the effect of tryptase did depend on an intact catalytic site because it was inhibited by leupeptin or benzamidine. Second, PAR-2 receptors were detected at the site of the HASM cell by means of immunocytochemistry, as recently reported by Hauck et al. (15). Third, trypsin, which, like tryptase, activates PAR-2 (25), induced a [Ca2+]i response similar to that induced by tryptase. Fourth, the AP SLIGKV, which corresponds to the tethered ligand that is exposed following proteolysis of the extracellular NH2-terminal domain of PAR-2 (24), also produced a [Ca2+]i response similar to that induced by tryptase, whereas the RP was without effect. Fifth, homologous desensitization of PAR-2 was demonstrated with tryptase, trypsin, and the AP SLIGKV. Such desensitization is a characteristic feature of PARs related to their mechanism of activation, i.e., enzymatic cleavage of the extracellular NH2-terminal domain. As a consequence, there is no PAR subtype cross-desensitization (34). Finally, PAR-2 desensitization seems to involve PKC because a specific inhibitor (calphostin C) magnified PAR-2 signaling, i.e., [Ca2+]i response induced by tryptase, trypsin, or AP. These results are in agreement with those by Bohm et al. (5), demonstrating that activation of PKC inhibits PAR-2 activation, whereas PKC inhibition increases this response in transfected epithelial cells.

Signal transduction mechanisms implicated in the tryptase-induced [Ca2+]i response appeared similar to those activated by the conventional agonists (ACh and histamine). That is, the Ca2+ source implicated in the PAR-2-mediated [Ca2+]i rise was primarily the intracellular Ca2+ store (presumably the sarcoplasmic reticulum), because TG, a sarcoplasmic Ca2+-ATPase inhibitor (13), completely inhibited tryptase-induced Ca2+ response, whereas Ca2+ removal, in the additional presence of EGTA, did not alter the tryptase-induced [Ca2+]i rise. The link between PAR-2 activation and intracellular Ca2+ release implicates PLC, because the tryptase-induced [Ca2+]i response was inhibited following pretreatment of cells with neomycin or U-73122, potent inhibitors of PLC (4, 8, 34), whereas it was unaltered by the negative control U-73343 (4, 34). As expected for G-protein-coupled receptors targeting PLC, the tryptase-induced response was not pertussis toxin sensitive (20, 27). These results strongly suggest that tryptase produces a [Ca2+]i release from an internal store via the IP3 receptor, as ACh (17, 23). This hypothesis is strengthened by the fact that a specific IP3-receptor antagonist, xestospongin C, significantly reduced the PAR-2-, trypsin-, and tryptase-induced Ca2+ responses. In this respect, the present results are in agreement with those obtained in another cell type expressing the PAR-2, the keratinocytes (34). It is also interesting to note that similar transduction mechanisms were previously reported in HASM cells in response to another protease, alpha -thrombin, which activates PAR-1, a different subtype of PAR (27).

A striking difference between the Ca2+ response induced by tryptase or alternative PAR-2 agonists and that by the ACh or histamine was the duration between the beginning of the ejection of the agonist and the peak of the response. This duration was approximately twice as long on PAR-2 activation and is likely to be related to the unique process of protease-induced receptor activation, which includes recognition of the receptor by the enzyme, cleavage of the receptor at a specific enzyme site, exposure of the tethered ligand, and, finally, binding of the tethered ligand.

Agonist-induced Ca2+ signaling in smooth muscle cells is often a complex temporal [Ca2+]i signal composed of a series of cyclic increases in [Ca2+]i, so-called Ca2+ oscillations (33). At the whole cell level, [Ca2+]i oscillations in airway smooth muscle cells are primarily IP3 dependent, involving a cyclic Ca2+ release-Ca2+ reuptake by intracellular store (31, 33). Because tryptase activates the IP3 pathway in HASM cells, its potential ability to produce Ca2+ oscillations deserves further discussion. In the present study, tryptase did not produce the so-called baseline-spiking Ca2+ oscillations, i.e., oscillations characterized by the [Ca2+]i returning to its resting value between each increase. In most of the cells, however, high concentrations of tryptase produced a Ca2+ response that evoked sinusoidal types of oscillations characterized by the [Ca2+]i remaining above its resting value between each increase (Figs. 2 and 4). The fact that, as discussed above, the rate of tryptase-induced [Ca2+]i rise was low may account for this phenomenon, because this rate plays a role in the control of the functioning of the IP3-receptor channel (21, 41). The use of real-time confocal imaging has provided new information on the mechanisms underlying Ca2+ oscillations at both the subcellular and whole cell level (19, 26, 28, 29, 38). Agonist-induced propagating oscillations of [Ca2+]i have been described in airway smooth muscle cells as the result of a repetitive release of Ca2+ through ryanodine-receptor channels following an initial IP3-induced Ca2+ release (19, 38). Spontaneous, localized Ca2+ transients (Ca2+ sparks), which represent elementary Ca2+ release through ryanodine-receptor channels, have been described in airway smooth muscle cells (38), and it has been demonstrated that agonist-induced [Ca2+]i oscillations represent a spatial and temporal integration of Ca2+ sparks (26). Therefore, the subcellular effect of tryptase on both Ca2+ sparks and spatial/temporal integration of such sparks should now be examined, taking into account that the tryptase-induced [Ca2+]i rise is delayed compared with more conventional agonists such as ACh.

This observation that tryptase is a potent activator of Ca2+ signaling at the site of airway smooth muscle has important implications in applied physiology and pathophysiology. The range of concentrations of tryptase that triggered a Ca2+ response in the present study (7.5-30 mU/ml) is similar to that inducing a contractile response in dog airways (37) or potentiating human isolated bronchi contractility (2). To further correlate the present data obtained in isolated cells with those in airways both in vitro and in vivo, it is noteworthy that the range of concentrations of the AP that triggered a Ca2+ response in the present study is also similar to that inducing a bronchomotor response in the live guinea pig (30). Although the physiological concentration of tryptase is difficult to estimate in airway tissue, the large number of mast cells in lung tissue (3) and the large quantities of tryptase present in these cells (36) suggest that such tryptase concentrations will be attained in vivo following mast cell activation. Moreover, there is evidence that the number of mast cells at the site of the airway smooth muscle increases under pathophysiological conditions (1, 2). In conclusion, this study provides evidence that tryptase, which is increasingly recognized as an important mediator in airway inflammation and hyperresponsiveness, is also a potent direct agonist at the site of airway smooth muscle. In our hands, the overall effect of tryptase on the mechanical activity of isolated HASM was very variable. In some rings, tryptase induced a large contraction; in some others, it produced relaxation; and in most of the rings it did not change the tone. This variability in the effect of tryptase may be because the integrative effect of tryptase is a combination of several opposite unitary effects. In this connection, it is noteworthy that relaxation of main bronchi in vitro was reversed to contraction by removal of epithelium in the guinea pig (30). Therefore, an understanding of the effect of lung tryptase requires understanding its relative role in both epithelial and smooth muscle cells.


    ACKNOWLEDGEMENTS

This study was supported by grants from Mutuelle Générale de l'Education Nationale and Conseil Régional d'Aquitaine (no. 980301115).


    FOOTNOTES

Address for reprint requests and other correspondence: R. Marthan, Laboratoire de Physiologie Cellulaire Respiratoire, Inserm E9937, Université Bordeaux 2, 146 rue Léo Saignat, 33076 Bordeaux Cedex, France (E-mail: roger.marthan{at}lpcr.u-bordeaux2.fr).

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.

Received 2 March 2001; accepted in final form 27 April 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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J APPL PHYSIOL 91(2):995-1003
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