Abstract
Neutrophils are infiltrated in airways of individuals with more severe and chronic asthma, with uncertain significance. Airway smooth muscle (ASM), apart from its contractile properties, is critically involved in the pathogenesis of asthma by producing inflammatory mediators. In the present study, we investigated the impact of neutrophil-derived elastase (NE) on ASM in terms of TGF-β1 release, and we explored the underlying mechanisms. Primary ASM cells were serum starved for 24 h before stimulation with NE (0.01–0.5 μg/ml). TGF-β1 in supernatant was determined by ELISA and mRNA quantified by real-time RT-QPCR. NF-κB nuclear translocation and activation was examined by Western blotting and κB-2 dEGFP reporter gene assay. Association of IL-1 receptor–associated kinase (IRAK) with MyD88 was studied by co-immunoprecipitation and Toll-like receptor 4 (TLR4) determined by FACS scan and Western blotting. We demonstrated that NE enhanced TGF-β1 release in a time-dependent manner. This induction was inhibited by actinomycin D (5 mM), cycloheximide (5 mM), and NF-κB inhibitors, including pyrrolidine dithiocarbamate (PDTC, 1 mM), aspirin (2.5 mM), and sodium salyicylate (2.5 mM). Stimulation with NE was rapidly followed by association of IRAK with MyD88, phosphorylation of IκBα, and nuclear translocation of p65 with increased transactivation activity. We also found that TLR4 levels were reduced upon NE treatment. These data suggest that NE upregulates TGF-β1 gene expression and release via My88/IRAK/NF-κB pathway, possibly through activation of TLR4, and shed light on a potential role of neutrophils in the pathogenesis of asthma.
Asthma is a disease of chronic airway inflammation, which is unique in that the airway wall is infiltrated by T lymphocytes, eosinophils, macrophages/monocytes, mast cells, and sometimes by neutrophils (1). In parallel with the chronic inflammatory process, airways remodeling characterized by an increase in muscle mass, mucous glands, and vessel area, as well as thickening of the reticular basement membrane, is almost always present in biopsies of individuals with asthma (2). Among these individuals, airway smooth muscle (ASM) is subject to a dominant pathologic change. In addition to contractile properties, ASM may contribute to airway inflammation of asthma by expression and secretion of proinflammatory cytokines and mediators, such as eotoxin (3) and IL-5 (4). In asthmatic airway remodeling, the mechanism of phenotype change in ASM has not yet been clear.
Neutrophils have been recovered in the airways of individuals with asthma, although their role in the pathogenesis of asthma remains unclear. The number of airway neutrophil increases during the late-phase response after an allergen challenge (5), in some patients who died within hours after an asthma exacerbation (6), and in some patients with long-standing asthma (7). Neutrophil-derived elastase (NE) is a serine protease involved in host defense against bacterial pathogens (8). NE is capable of hydrolyzing a broad spectrum of extracellular matrix (ECM) and cellular proteins, such as elastin, interstitial collagens, proteoglycans, fibronectin, laminin, and others (9). Increasing evidence also indicates that NE can provoke a variety of cellular responses. For instance, NE has been identified to induce the expression of IL-8 (10) and secretory leukocyte protease inhibitor (SLPI) (11) in epithelial cells, and to induce ASM cell proliferation (12).
Transforming growth factor-β (TGF-β), made up of three isoforms (TGF-β1, -2, and -3), is a multifunctional cytokine contributing to the initiation and resolution of inflammatory events, impacting on the cell growth, and playing an important role in tissue repair and fibrosis (13). Compared with normal subjects, TGF-β concentrations are higher in bronchoalveolar lavage fluid of individuals with asthma (14). TGF-β gene expression is upregulated in bronchial tissue from patients with severe asthma (15) and has been demonstrated to correlate with the thickness of subepithelial basement membrane. Thus, TGF-β may contribute to the pathogenesis of airway remodeling in asthma (16).
NF-κB is a transcription factor of particular importance in immune and inflammatory responses (17). The activated form of NF-κB is a heterodimer consisting of two proteins, a p65 subunit and a p50 subunit, or other subunits (rel, rel-B, p52, v-rel) (18). In unstimulated cells, NF-κB is found in cytoplasm bound to IκBs. Stimulation with cytokines, reactive oxygen species, and microorganisms induces phosphorylation and degradation of IκBs, followed by translocation of NF-κB to the nucleus, binding to its response elements in promoters and leading to transcriptional initiation of responsive genes (19).
In the present study, we have examined the role of NE as an inducer of cytokine production by ASM. We demonstrated that NE enhanced the release of TGF-β1 in primary cultures of human ASM cells. By using quantitative real-time reverse transcriptase–polymerase chain reaction (RT-QPCR), we also revealed an increase in mRNA level and provided evidence that a transcriptional and translational mechanism was involved. Furthermore, the NE-induced ASM TGF-β1 secretion was preceded by activation of NF-κB and was blocked by pretreatment with NF-κB inhibitors. Finally, we showed a rapid association of MyD88 and IL-1 receptor–associated kinase (IRAK) after NE stimulation. These findings demonstrate that NE activates the MyD88/IRAK/NF-κB pathway in generation of TGF-β1 from ASM.
The human bronchial smooth muscle cells (hBSMC; Clonetics Corporation, San Diego, CA) were cultured in 25 cm2 flasks, in culture medium for hBSMCs supplemented with 10% FBS (fetal bovine serum), human epidermal growth factor (hEGF, 0.5 g/ml), insulin (5 mg/ml), human fibroblast growth factor (hFGF, 1 μg/ml), gentamicin (50 μg/ml), and amphotericin B (0.05 μg/ml) (SmGM-2 BulletKit; Clonetics) at 37°C, 5% CO2. When the cells became confluent, they were subcultured in the same medium. At confluence, cells obtained from 25 cm2 flasks were passaged using 0.25% trypsin in 0.02% EDTA. Medium was changed every other day. Cells before confluence at passages 3 to 7 were detached from culture flasks with 0.25% trypsin in 0.02% EDTA and were used for later experiments. Confluent cells were growth arrested in serum-free DMEM for 24 h before experiments. ASM cell number counted by a hemocytometer and DNA synthesis (12) did not increase 6–12 h after stimulation with human NE (0.01, 0.05, 0.1, and 0.5 μg/ml). The cellular viability was determined by MTT assay (Roche Diagnostics, Basel, Switzerland) according to the manufacturer's instructions.
Human NE, actinomycin D, cycloheximide, pyrrolidine dithiocarbamate (PDTC), aspirin (acetylsalicylic acid), and sodium salicylate (NaSal) were purchased from Sigma Chemical Co. (St. Louis, MO). Elastase inhibitor II and a protease inhibitor cocktail (500 μM AEBSF, HCl, 150 nM aprotinin, 1 μM E-64, 0.5 mM EDTA, disodium salt, and 1 μM leupeptin hemisulfate) were obtained from Calbiochem (La Jolla, CA). Monoclonal neutralizing antibodies against human TLR4, IL-1α, and IL-1β were obtained from R&D Systems (Minneapolis, MN). Monoclonal antibodies against proteinase-activated receptor-2 (PAR2) were obtained from Bachem Bioscience, Inc. (King of Prussia, PA).
To investigate the effects of chromogenic substrate for NE, NE at final concentrations of 0.01–1 μg/ml was added to assay buffer (0.45M Tri-Base, 2M NaCl, pH 8.0) and elastase substrate I (MeOSuc-Ala-Pro-Val-pNA; Calbiochem) for 5 min. The cleavage was monitored at 410 nm spectrophotometer, and the slope for NE activity was determined.
The concentrations of TGF-β1 in supernatants were measured with specific ELISA kits (R&D Systems) using the quantitative immunometric sandwich enzyme immunoassay technique. For the assay, the frozen supernatants prepared from cultured ASM were thawed at room temperature and were first activated by being incubated with 1 N HCl for 10 min and neutralized by 1.2 N NaOH/0.5 M N-2-hydroxyethylpiperazine-N′-ethane sulfonic acid. Activated samples were then transferred to the wells of rigid, flat-bottom microtiter plates coated with TGF-β1 soluble receptor Type II. After incubation and thorough washing, horseradish peroxidase (HRP)-conjugated antibodies directed against TGF-β1 were added to the test wells. After a second incubation, excess HRP-conjugated antibody was removed by washing. The HRP substrate was then added and the color intensity was measured with a microtiter plate reader.
Total RNA was extracted from 8 × 105 ASM cells and cultured for 0.5–4 h in the presence or absence of NE, using the guanidine thiocyanate/phenol/chloroform method. cDNA was reverse-transcribed from isolated RNA by incubating 1.5 μg of DNase-treated RNA with a first-strand synthesis Kit from Advanced Biotechnologies Ltd (Surrey, UK). cDNA amplifications were done by quantitative PCR using the light cycler and the double-stranded DNA binding dye SYBR Green 1 (Roche Diagnostics). PCR mixtures contained 0.5 μM of either glyceraldehyde-3-phosphate dehydrogenase (GAPDH) or TGF-β1–specific primers. The samples were denatured at 95°C for 10 min followed by 45 cycles of annealing and extension at 95°C for 15 s, 55°C for 5 s, and 72°C for 10 s. The melting curves were obtained at the end of amplification by cooling the samples to 65°C for 15 s followed by further cooling to 40°C for 30 s. Data were analyzed by standard curve method of absolute quantification using the LightCycler analysis software with serial dilutions of known quantities of GAPDH and TGF-β1 cDNA PCR products. Values were normalized to GAPDH expression. PCR products were further confirmed by gel electrophoresis and melt curve analysis. The TGF-β1 primers included 5′ GTG GAA ACC CAC AAC GAA A 3′ (sense) and 5′ TAA GGC GAA AGC CCT CAA T 3′ (antisense). GAPDH primers were obtained from GIBCO-BRL (Gaithersburg, MD).
Cytoplasmic and nuclear extracts were subjected to SDS-polyacrylamide gel electrophoresis and blotted onto nitrocellulose filters. NF-κB subunit p65 was detected with a mouse anti-human p65 antibody (Transduction Laboratories) and an alkaline phosphatase–conjugated anti-mouse antibody. Phosphorylated IκB was detected with a rabbit anti-human phosphor-IκBα antibody (Cell Signaling Technology Inc., Danvers, MA) and an alkaline phosphatase–conjugated anti-rabbit antibody.
TLR4 protein was detected on total protein of ASM cells after exposure to NE (0.5 μg/ml) for 20 min with a mouse IgG2A anti-human TLR4 antibody (Serotec, Oxford, UK) and an alkaline phosphatase-conjugated anti-rabbit antibody. Blots were developed by adding of 5-bromo-4-chloro-3-indole phosphate/nitroblue tetrazolium solution (Sigma) and were then exposed to XAR-5 film.
To quantify TLR4 and PAR2 cell surface expression, ASM cells were seeded in 8-well chamber slides in serum-free medium for 24 h before exposure to NE (0.5 μg/ml, 20 min). Cells were labeled with mouse IgG2A anti-human TLR4, PAR2, and a fluorescein isothiocyanate (FITC)-labeled anti-mouse F(ab)2 (Dako). Isotype control samples were also prepared. The cells were extensively washed twice and the expression of TLR4 or PAR2 was measured by FACScan flow cytometer equipped with an argon ion laser (Becton Dickinson, Mountain View, CA). Off-line analysis was performed using QUEST software as supplied by Becton Dickinson. TLR4 or PAR2 expression was expressed as the mean fluorescence intensity.
Confluent ASM cells in 6-well dishes were transiently transfected in Optimem (Life Technologies, Inc., Gaithersburg, MD) with 10 μl of LipofectAMINE2000 (Life Technologies, Inc.), 4 μg of pNF-κB–d2EGFP, or pEGFP-C2 vector (BD Biosciences Clontech, Palo Alto, CA), used to determine transfection efficiency. The cells were added with serum-free DMEM-F12 5 h later. Eighteen hours after transfection, the media were replaced with fresh serum-free DMEM-F12 and the cells were treated with 0.5 μg/ml NE or let untreated for 4 h. The cells were examined with a fluorescence microscope. In parallel experiments the cells were also harvested with 0.25% trypsin in 0.02% EDTA for flow cytometric analysis.
ASM cells plated on 10-cm dishes were treated with 0.5 μg/ml NE for variable times (from 0–10 min). The cells were than harvested, lysed in 200 μl of PD buffer (40 mM Tris-HCl, pH 8.0, 500 mM Nacl, 0.1% Nonidet P-40, 6 mM EGTA, 10 mM β-glycerophosphate, 10 mM NaF, 300 μM sodium orthovanadate, 2 mM PMSF, 10 μl/ml aprotinin, 1 μg/ml leupeptin, and 1 mM DTT), and centrifuged. The supernatant was then immunoprecipitated with a polyclonal antibody against MyD88 (eBioscience, San Diego, CA) in the presence of A/G-agarose beads overnight at 4°C. The immunoprecipitated beads were then washed three times with PD buffer. The sample were fractionated on 7.5% SDS-PAGE, transferred to nitrocellulose membrane, and subjected to immunoblot analysis with human IRAK (Calbiochem) and MyD88 antibodies.
Data were analyzed with the GraphPad Prism 3.0 software package (GraphPad Software, San Diego, CA). Results are expressed as mean ± SE. Unpaired comparison was analyzed by Mann-Whitney U test. In the time course study of TGF-β1 mRNA induction, one-way ANOVA with Dunnet's multiple comparison post-test was used to validate the trend. Differences were considered significant when the P value was ⩽ 0.05.
To determine the effect of NE on TGF-β1 release, confluent, growth-arrested ASM cells were treated with different concentrations of NE (0.01, 0.05, 0.1, and 0.5 μg/ml) or were left untreated for 6 h. NE at the concentration of 0.5 μg/ml significantly increased TGF-β1 concentration (316.2 ± 68.8 pg/ml) compared with the basal level (58.5 ± 19.0 pg/ml, n = 5, P < 0.01) (Figure 1A). Exposure of ASM cells to higher concentrations of NE (1 or 5 μg/ml) were excluded from the present study due to a high magnitude of cellular toxicity (determined by MTT assay). Although the effect of NE on TGF-β1 production was not dose-dependent, the purity assay of NE activity showed a dose-dependent cleavage effect on chromogenic substrate (data not shown). ELISA assay for TGF-β1 protein detects both active and nonactive forms of TGF-β1. However, no spontaneous active form was detectable without pre-acidification of the culture supernatants in the presence or absence of NE. Incubation of the supernatant of untreated ASM cells with NE (0.5 μg/ml) for 6 h did not affect the basal level of TGF-β1 (50.3 ± 18.7, n = 4) when compared with those in the absence of NE (51.0 ± 20.3, n = 4).
Figure 1. Effect of NE on TGF-β1 release. (A) ASM cells were stimulated with (0.01–0.5 μg/ml, n = 5) for 6 h in the presence or absence of neutrophil inhibitor II (EI II, 100 μg/ml, n = 5) or protease inhibitor cocktail (PI, 500 μM). Supernatants were collected and assayed by TGF-β1 ELISA. Data are mean ± SE. *P < 0.01 compared with the control group (n = 5); #P < 0.01 compared with 0.5 μg/ml of NE. (B) Time course of NE-stimulated TGF-β1 release. ASM cells were treated with (open circles, n = 5) or without (closed circles, n = 5) 0.5 μg/ml NE for 0, 6, and 24 h. Data are mean ± SE. *P < 0.01 compared with the corresponding time controls; #P < 0.01 compared with the same treatment at 6 h. (C) MTT assay for ASM cells viability after exposure to NE (0.05–1.0 μg/ml, n = 5) for 6 h. Cell viability decreases at 1.0 μg/ml NE.
[More] [Minimize]To determine whether the enhanced TGF-β1 secretion is due to the NE-specific activity, NE was used in the presence of a potent irreversible inhibitor of human NE, elastase inhibitor II (EI II), or a protease inhibitor cocktail (PI). Either pretreatment with EI II (100 μg/ml) or a protease inhibitor cocktail resulted in complete inhibition of the NE-mediated release of TGF-β1.
The time course of TGF-β1 secretion from ASM cells is shown in Figure 1B. In the absence of NE, there was a constitutive TGF-β1 production increasing progressively from 6 to 12 to 24 h (58.2 ± 19.0 pg/ml, 87.3 ± 56.1 pg/ml, and 158.1 ± 80.8 pg/ml respectively, n = 5, P < 0.01). NE increased TGF-β1 production significantly at 6 h (166.4 ± 36.2 pg/ml) and continued to enhance the production to 288.5 ± 64.3 pg/ml at 12 h and 494.2 ± 73.7 pg/ml at 24 h (P < 0.01 compared with control ASM cells at comparative time points). Since there was a significant increase in cell number after incubation with NE for 24 h, longer time culture was not further examined.
ASM cells were pretreated for 30 min with actinomycin D (5 mM), a transcription inhibitor, or cycloheximide (5 mM), a translation inhibitor, before NE treatment or without treatment. The NE-induced TGF-β1 release in the supernatant (from 58.2 ± 14.5 pg/ml of a basal level to 148.8 ± 21.6 pg/ml, P < 0.01) was inhibited either by actinomycin D (92.0 ± 26.1 pg/ml, P < 0.01) or cycloheximide (87.0 ± 36.4 pg/ml, P < 0.01) (Figure 2A). These data suggest that the NE-induced TGF-β1 release involves both transcription and translation mechanisms.
Figure 2. Effect of NE on TGF-β1 transcription. (A) Effect of actinomycin D (ACD) and cycloheximide (CHX) on NE-induced TGF-β1 secretion. ASM cells were incubated for 6 h with or without NE (0.5 μg/ml) after pretreatment with actinomycin D (5 mM), cycloheximide (5 mM), or without pretreatment for 30 min (n = 5). Data are mean ± SE. *P < 0.01 compared with control; #P < 0.01 compared with NE. (B) Effect of NE on the induction of TGF-β1 mRNA expression. Total RNA was extracted from untreated cells or cells (4 × 105 cells/ml) treated with NE (0.5 μg/ml, n = 4) after 0.5–6 h of culture and 1.5 μg was reverse-transcribed into cDNA and used as template in quantitative LightCycler PCRs. Values are expressed as fold induction of TGF-β1 mRNA from basal level after normalization to GAPDH. Data are mean ± SE. *P < 0.01 compared with time 0.
[More] [Minimize]To further confirm an increase in transcription, TGF-β1 mRNA was measured by RT-QPCR. Figure 2B demonstrates that NE time-dependently induced TGF-β1 mRNA accumulation, with a maximal and significant induction of 2.3 ± 0.7-fold at 4 h (P < 0.01 by one-way ANOVA with Dunnett's multiple comparison test) during the study period.
NE induces NF-κB activation in bronchial epithelial cells (20). We therefore explored which signal pathways mediate the NE-induced TGF-β1 gene upregulation. ASM cells were left untreated or stimulated with 0.5 μg/ml NE with or without pretreatment by pyrrolidine dithiocarbamate (PDTC, 1 mM), a nonspecific NF-κB inhibitor, aspirin (Asp, 2.5 mM), or sodium salicylate (NaSal, 2.5 mM), both inhibitors of IκB kinase-β (21), for 30 min. As shown in Figure 3, pre-incubation with PDTC, aspirin, or NaSal abolished the NE-induced TGF-β1 production (144.7 ± 31.2 pg/ml, n = 5, P < 0.01; 210.4 ± 46.3 pg/ml, n = 5, P < 0.05; 144.8 ± 10.4 pg/ml, n = 5, P < 0.01, respectively), suggesting NF-κB was mediated in NE-induced TGF-β1 secretion.
Figure 3. Inhibition of NE-induced TGF-β1 expression by NF-κB inhibitors. Cells were untreated or stimulated with 0.5 μg/ml NE for 6 h, with or without pretreatment with pyrrolidine dithiocarbamate (PDTC, 1 mM), aspirin (Asp, 2.5 mM), or sodium salicylate (NaSal, 2.5 mM) for 30 min. Data are mean ± SE. #P < 0.01 compared with the control group (n = 5); **P < 0.01 compared with 0.5 μg/ml of NE (n = 5); P < 0.05 compared with 0.5 μg/ml of NE (n = 5).
[More] [Minimize]Upon NE stimulation (0.5 μg/ml), phosphorylation of IκB-α rapidly occurred at 5 min (Figure 4A), followed by an increase in nuclear p65 by 2.5 ± 0.9-fold (Figure 4B, P < 0.05, n = 3) at 10 min and returned to the basal levels at 30 min. These data demonstrate that p65 was released from the cytoplasm and translocated to the nucleus by NE stimulation.
Figure 4. Effect of NE on NF-κB activation. ASM cells were treated with 0.5 μg/ml NE for 0–30 min Cytoplasmic extracts were assessed for phosphorylated IκBα (A) and the nuclear extracts were assessed for p65 (B, upper panel) by Western blot analysis. Data are representative of three independent experiments with similar results. Bar graphs show quantitative analysis of scanning densitometric values of nuclear p65 (B, lower panel). Data are mean ± SE. *P < 0.05 compared with basal levels (n = 3).
[More] [Minimize]To directly determine NF-κB activation, we used reporter gene assays. ASM cells were transiently transfected with pNF-κB-d2EGFP. Eighteen hours after transfection, the cells were treated with 0.5 μg/ml NE for 4 h, and the fluorescence intensity of the transfected cells was examined by fluorescence microscopy. Transfection efficiency was determined by transfection with pEGFP-C2. As shown in Figures 5A–5D, NE-stimulated cells appeared strong d2EGPF fluorescence intensity, suggesting an augmentation in NF-κB transactivation activity. To further quantify this activity, we used flow cytometry to measure fluorescence intensity. We found a 1.8 ± 0.1-fold increase in mean fluorescence intensity (MFI) in the NE-treated cells compared with unstimulated cells (Figure 5E, n = 5, P < 0.01). These results confirm the observation of NF-κB activation in Western blotting.
Figure 5. Induction of reporter genes. ASM cells were transiently transfected with pNF-κB–d2EGFP. Eighteen hours after transfection, cells were treated with 0.5 μg/ml NE or left untreated for 4 h. The cells were collected for light microscopy (A, control; C, elastase; magnification: ×200) and fluorescence microscopy examinations (B, control; D, elastase; magnification: ×200) on cytospin and for flow cytometry analysis (E). Values are expressed as fold induction of MFI (mean fluorescent intensity) from control. Data are mean ± SE. *P < 0.01 compared with control (n = 5).
[More] [Minimize]MyD88/IRAK/TRAF pathway, an upstream NF-κB signal pathway, has been proved to mediate the NE-induced IL-8 upregulation in human bronchial epithelium (20), we next examined whether NE stimulates IRAK activation in ASM by using co-IP. Upon stimulation with NE (0.5 μg/ml), IRAK was rapidly associated with MyD88 as early as 5 min and partially sustained at 10 min (Figure 6). This kinetic is temporally in agreement with that of IκB-α phorylation and NF-κB activation and suggests that MyD88/IRAK pathway is the upstream pathway of NF-κB–mediated TGF-β1 gene expression by NE stimulation.
Figure 6. NE induced activation of MyD88 and IRAK. ASM cells were treated with 0.5 μg/ml NE for 0–10 min. Cell extracts were then immunoprecipitated with anti-MyD88 antibody and probed with antibodies against IRAK and MyD88 after Western blot transfer. The experiment was repeated, with a similar result. IP, immunoprecipitation; IB, immunoblot.
[More] [Minimize]IRAK was first described as a signal transducer for the proinflammatory cytokine IL-1 and was later implicated in signal transduction of other members of the Toll-like receptor (TLR)/IL-1 receptor (IL-1R) family (22). We next asked whether IL-1 and IL-1R are involved in the upregulation of TGF-β1 production by NE. ASM cells were pretreated with or without neutralizing antibodies against IL-1α (0.1 or 1 μg/ml), IL-1β (0.1 or 1 μg/ml), IL-1RI (0.1, 1, 10 μg/ml), and IL-1RII (0.1, 1, 10 μg/ml) for 30 min, followed by NE stimulation (0.5 μg/ml) or left untreated for 6 h. By ELISA, we failed to find any inhibition effect on NE-induced TGF-β1 release with these neutralizing antibodies. In addition, there was no detectable level of IL-1α, or IL-1β in the supernatants of NE-stimulated ASM (data not shown). These results exclude the involvement of IL-1 and IL-1R in the NE induction of TGF-β1 secretion.
Furthermore, NE was shown to activate NF-κB through TLR4 (23), via IRAK/MyD88/TRAF6 pathway (20). In subsequent experiments, TLR4 protein expression on ASM cell surface was analyzed by flow cytometry, and was found decreased after NE (0.5 μg/ml) treatment (Figure 7A). Western blot analysis of TLR4 protein on ASM cells was also decreased with NE (0.5 μg/ml) (Figure 7B). Pretreatment with a blocking antibody to TLR4, but not PAR2, attenuated the effect of NE (0.5 μg/ml) on TGF-β1 production (Figure 7C) (Figure 8B). NE did not affect the expression of PAR2 on ASM cells (Figure 8A).
Figure 7. NE decreases expression of TLR4 protein. ASM cells were stimulated with 0.5 μg/ml NE for 20 min and the expression of TLR4 on ASM cell surface was analyzed by flow cytometry (A) and Western blot (B). Three identical experiments independently performed gave similar result. (C) ASM cells were pretreated with or without a neutralizing antibody to TLR4 (TLR4) before exposure to NE (0.5 μg/ml, elastase) for 6 h. Data are mean ± SE (n = 5 in each group). **P < 0.01 compared with corresponding control; #P < 0.05 compared with ASM cells treated with NE alone.
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Figure 8. NE-induced TGF-β1 release is not mediated via PAR2. ASM cells were stimulated with 0.5 μg/ml NE (elastase) for 20 min, and the expression of PAR2 on ASM cell surface was analyzed by flow cytometry (A). Three identical experiments independently performed gave similar result. (B) ASM cells were pretreated with or without a neutralizing antibody to PAR2 (+PAR2) before exposure to NE (0.5 μg/ml) for 6 h. Data are mean ± SE (n = 5 in each group). **P < 0.01 compared with control.
[More] [Minimize]This is the first study showing that NE enhances TGF-β1 gene expression and protein production in human ASM cells. Given that the NE-stimulated response was inhibited by actinomycin D or cycloheximide, the NE-induced TGF-β1 production involves de novo protein production. Upon NE stimulation, IκBα was rapidly phosphorylated, followed by p65 nuclear translocation. The fact that pretreatment with variable NF-κB inhibitors prevented the NE-induced TGF-β1 release further supports the role of NF-κB. NE increased association of IRAK with MyD88, which has been shown to be followed by induction of MyD88/IRAK/TRAF-6 (tumor necrosis factor receptor-associated factor 6) pathway, and in turn activation of NIK and NF-κB activation (24, 25).
NE has been reported to exhibit several functions in addition to its proteolytic activity, such as induction of IL-8 (10). In the present study, the NE-induced TGF-β1 production was maximal with NE at 0.5 μg/ml. Higher concentration of NE was not further examined due to a high magnitude of cellular toxicity. The dose-dependent cleavage effect on chromogenic substrate confirms the purity of NE used in this study, and stands against the possibility that impurity of NE results in a lack of dose–response effect of NE. TGF-β1 is secreted in latent forms and becomes active after cleavage by proteolytic enzymes (13). TGF-β1 was measured by using an ELISA assay which allows us to distinguish active and latent forms of the TGF-β1. The antibody only recognizes the active form. Thus, the concentrations of TGF-β1 measured before acidification represent the constitutively active form, and those measured after acidification that activated latent TGF-β1 represent the total TGF-β1 present in the samples. No spontaneous active form was detectable without pre-acidification of the culture supernatants in the presence or absence of NE, suggesting that the increased level of TGF-β1 after NE treatment was not attributed to proteolytic activity of NE by which the latent form of TGF-β1 is activated. It is also possible that NE may cleave the antigen of TGF-β1, resulting in an increase in antibody-binding affinity of ELISA. However, we incubated the supernatant of untreated ASM cells with NE (0.5 μg/ml) for 6 h, but did not find any change in the basal level of TGF-β1. With real-time RT-QPCR we have provided evidence that NE upregulates expression of TGF-β1 mRNA in ASM cells. In addition, the findings that pretreatments with actinomycin D or cycloheximide attenuate the response of ASM to NE further confirm the involvement of both transcriptional and translational mechanisms. These findings, together with the observation that elastase releases TGF-β1 from the extracellular matrix of human epithelial and endothelial cells (26) and from mouse lungs (27), suggest that elastase has the potential to upregulate TGF-β1 at multiple levels, transcriptioanlly in cells and post-translationally in extracellular storage sites.
In this article, we show that NF-κB is involved in the NE enhancement of TGF-β1 expression. NE-induced TGF-β1 release was inhibited by the NF-κB inhibitor PDTC, aspirin, and sodium salicylate, compounds with anti-IKKβ activity. By using Western blotting, we clearly demonstrated that NE induced IκBα phosphorylation and p65 nuclear translocation. Reporter gene assays further confirmed the activation of this transcription factor. Importantly, an NF-κB response element in the TGF-β1 promoter has recently been identified which was shown to mediate TGF-β1 transcription stimulated by IL-1β (26). Our data are also in agreement with the finding that treatment with relA antisense oligonucleotides inhibited TGF-β1 mRNA expression in fibrosarcoma cells (28) and monocytes from patients with idiopathic myelofibrosis (29). In the latter study, it was argued that IL-1α was released after NF-κB activation and that it then autostimulated the production of TGF-β1 (29). Our findings that there was no detectable IL-1α or IL-1β in culture supernatants after NE treatment, and that pretreatments of cells with blocking antibodies against IL-1α or IL-1β had no effect on TGF-β1 production, exclude the involvement of IL-1 cytokine in the response to NE. Experiments with IL-1RI and IL-1RII–neutralizing antibodies further exclude these receptors from targets of NE. Taken together, our data suggest that NE stimulate TGF-β1 expression through an NF-κB–mediated mechanism.
TLRs compose a large family with 10 members (TLR1–10) (30–32), which play an important role in the recognition of microbial components, with subsequent activation of innate immunity leading to development of adaptive immune responses (33). The prototypical TLR is TLR4, which is the recognized receptor for lipopolysaccharide (LPS) (34). To transmit its signal, TLRs recruit adaptor proteins such as MyD88 (myeloid differentiation factor 88) that leads to phosphorylation of IκB, allowing nuclear translocation of NF-κB. NE has been shown to stimulate IL-8 release from human bronchial epithelial cells through TLR4 (23) and the downstream IRAK/MyD88/TRAF6 pathway to activate NF-κB (20). In the present study, NE decreased TLR4 expression on ASM cells with increased expression of IRAK and MyD88, and then subsequent enhanced IRAK co-precipitation with MyD88, indicating activation and increased recruitment of IRAK to MyD88. Treatment with LPS was also found to decrease TLR4 surface expression on ASM cells in this study. NE was also reported to decrease TLR4 surface expression on bronchial epithelium to activate NF-κB (35). Thus, the mechanism by which NE decreased TLR4 surface expression may be similar to that of LPS-induced TLR4 internalization or alternatively could be a result of cleavage by NE (35, 36). Pretreatment with a TLR4-blocking antibody attenuates NE-induced TGF-β1 production, confirming that TLR4 is mediated in this response. PAR-2 has also been reported to mediate TGF-β1 synthesis at a post-transcriptional level through activation of the ERK MAP kinase cascade (37). In the present study, NE did not affect the surface expression of PAR2. Furthermore, the NE-stimulated TGF-β1 secretion was not affected by the specific ERK inhibitor PD98059 or a blocking antibody against PAR2, precluding the involvement of PAR2 in this response. Taken together, our present results indicate that NE activates IRAK/MyD88 through TLR4 in ASM cells to activate NF-κB pathways.
TGF-β1 is a potent profibrotic cytokine and has been suggested to play a role in the fibrotic changes occurring within asthmatic airways (16, 38). Although the predominant effect of TGF-β1 on cell proliferation is inhibitory, the mitogenic action of TGF-β1 has been demonstrated in ASM cells (39) and has been suggested to be associated with the hyperplastic nature of ASM cells in chronic asthma and bronchopulmonary dysplasia. These observations and our present study address a potential role of NE, by inducing TGF-β1 production from ASM cells, in airway fibrosis and ASM hyperplasia, both major characters of airway remodeling in asthma.
Endogenous NE inhibitors such as α1-antiprotease or SLPI are present in the lower respiratory tracts. Exploration of the inhibitory effect of endogenous broad spectrum inhibitors on NE-induced responses may be needed to reflect the actual role of NE in human pathophysiologic conditions.
In summary, our data reveal an effect of NE on TGF-β1 gene induction and protein production from human ASM cells. The effect appears to be proximally mediated by the activation of NF-κB, possible via the MyD88/IRAK pathway. Given the fact that there are abundant neutrophils in airways of more severe or chronic asthma and that TGF-β1 contributes to airway fibrosis and ASM hyperplasia in chronic asthma and in a variety of other lung conditions, our findings not only suggest a role of neutrophils in the pathogenesis of airway remodeling but also help identify potential therapeutic targets. This may be achieved by inhibiting NE itself or any signaling components identified in this study to be involved in NE-induced TGF-β1 expression.
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