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* Division of Respiratory Medicine, Respiratory and Stress Care Center, Kagoshima University Hospital, Sakuragaoka, Kagoshima, Japan; and Laboratory of Molecular Immunoregulation, Center for Cancer Research, National Cancer Institute, Frederick, MD 21702
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Abstract |
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, and matrix metalloproteinase-9 (MMP-9) from CD14-positive BALF cells in a p38 MAPK-dependent manner. In contrast, DDR1 activation of CD14-positive BALF cells in the other groups did not induce the production of these chemokines or MMP-9. These chemokines and MMP-9 contribute to the development of IPF and, therefore, we suggest that DDR1 might be associated with the pathogenesis of IPF in the tissue microenvironment. |
Introduction |
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The basement membrane is a specialized form of the extracellular matrix (ECM), and a complex of collagen occupies a substantial portion of the basement membrane structure in the lung. The ECM is critically important in maintaining the structural integrity of the alveolar wall. In the development of IPF, disruption of the epithelial basement membrane is associated with inflammatory cell activation, including alveolar macrophages, and this interaction is considered to be the key event leading to intraluminal fibrosis (8, 9). It is also known that the amount of collagen increases during this process (10). Thus, the interaction between inflammatory cells and ECM is probably associated with the development of IPF; however, information regarding the molecular mechanisms is limited.
Discoidin domain receptor 1 (DDR1) is a receptor tyrosine kinase activated by binding with its ligand, collagen (11, 12). DDR1 has a unique extracellular domain that is homologous to discoidin 1 of Dictyostelium discoideum (13). DDR1 is constitutively expressed in normal tissues, such as lung, kidney, colon, and brain, and in tumor cells of epithelial origins such as mammary, ovarian, and lung carcinomas (13). Five DDR1 isoforms (a, b, c, d, and e) can be generated by alternative splicing of the DDR1 gene (13, 14). We previously reported that the expression of two DDR1 isoforms, DDR1a and DDR1b, could be induced in vitro in human leukocytes, including neutrophils, monocytes, and lymphocytes. In vivo, studies showed that tissue-infiltrating mononuclear cells, particularly macrophages, were positive for DDR1 mRNA (15). The DDR1a and DDR1b isoforms differ from each other by an in-frame insertion of 111 bp that codes for an additional 37-aa peptide in the proline-rich juxtamembrane region. The 37-aa insertion in DDR1b contains the LXNPXY motif that corresponds to the consensus-binding motif of the Shc phosphotyrosine-binding domain (13). In addition, we recently found that the activation of DDR1 isoforms, most likely DDR1b, up-regulated the production of chemokines in macrophages in a p38 MAPK-dependent manner (16), including MCP-1, which plays an important role in the pathogenesis of IPF (17). Autophosphorylation of DDR1b was highly detectable as early as 30 min after DDR1 activation, peaked at 60 min, and returned to basal levels by 120 min (16, 18, 19). In contrast, autophosphorylation of DDR1a was first detected 90 min after DDR1 activation, peaked at 4 h, and became undetectable by 12 h (19). Furthermore, DDR1 is reported to be associated with matrix metalloproteinase-9 (MMP-9) production (20, 21), which plays a pivotal role in the lung remodeling of IPF (22).
The DDR1 ligand, collagen (10), is abundant in IPF, and human monocyte-derived macrophages can express endogenous DDR1 in vitro (15, 19). Therefore, we hypothesized that the alveolar macrophages of IPF patients might express functional DDR1. We investigated DDR1 expression and chemokine production from CD14-positive bronchoalveolar lavage fluid (BALF) cells, which are considered to be alveolar macrophages, by the activation of DDR1. We also examined DDR1 signaling in CD14-positive BALF cells obtained from IPF patients. We found that the CD14-positive BALF cells of IPF patients expressed DDR1, with the DDR1b isoform being the predominant form, and that DDR1 activation induced its autophosphorylation, followed by the recruitment of the DDR1b adaptor protein Shc. DDR1 activation also induced chemokine and MMP-9 production via a p38 MAPK-dependent pathway.
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Materials and Methods |
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This study was reviewed and approved by the Kagoshima University Faculty of Medicine Committee on Human Research. We investigated 28 IPF patients (mean age = 56.4 ± 12.3 years old, male:female = 20:8), 13 chronic obstructive pulmonary disease (COPD) patients (mean age = 57.2 ± 14.3 years old, male:female = 8:6), and 14 healthy volunteers (mean age = 57.8 ± 17.8 years old, male:female = 6:8). All patients and volunteers provided written informed consent to participate in the study.
The diagnosis of IPF was based on previously published international criteria for idiopathic interstitial pneumonias (1). The diagnosis of COPD was based on the Global Initiative for Chronic Obstructive Lung Disease guidelines for COPD (23).
A bronchofibrescope (Olympus BF type p20; Olympus) was wedged into the right B4 segment of the lung to collect BALF cells. Four 40-ml aliquots of sterile physiological saline were instilled at 37°C, and recovered by gentle suction. The recovered fluid was immediately filtered through sterilized gauze, and the lavage fluid was centrifuged in a cytometer (KN-70; Kubota) at 44 x g for 5 min and stained with May-Giemsa stain to identify cell populations. Five hundred cells, excluding epithelial cells, were identified per slide to establish differential cell counts, and the counts were expressed in percentages. Concurrently, 1 x 105 cells were suspended in 50 µl of cold PBS containing 0.1% sodium azide, 10 ng/ml BSA, and 20 µg/ml human IgG and incubated for 10 min on ice. The cells were then incubated for an additional 15 min on ice with FITC-conjugated CD14 mAb (M5E2; BD Biosciences) and mouse anti-human CD29 mAb (MAR4, 
1 integrin; BD Biosciences) or anti-human DDR1 mouse IgG mAb (48B3; Santa Cruz Biotechnology). The cells were washed with PBS and incubated with biotin-conjugated goat anti-mouse IgG Ab for 15 min on ice. Cells were again washed with PBS and incubated with PE-conjugated streptavidin for 15 min on ice. At the end of the incubation period, 7-aminoactinomycin D (BD Pharmingen) was added to each tube. The cells were washed with PBS, and subsequently analyzed by flow cytometry using a FACScan (BD Biosciences). Dead cells, identified by the 7-aminoactinomycin D incorporation, were gated out. Results were processed using the CellQuest software (BD Biosciences), as described previously (24). BALF fluids were stored at –20°C for further analysis.
CD14-positive cells in BALF from each group were selected using magnetic beads (Miltenyi Biotec), according to the manufacturer’s protocol, and used for further analysis. Following selection, the CD14-positive cells were also stained with May-Giemsa stain to identify cell populations.
Immunohistochemistry
Biopsied lung tissues obtained from three IPF patients were examined by immunohistochemical staining for DDR1 using a rabbit anti-DDR1 Ab (Santa Cruz Biotechnology) or anti-human CD14 mouse IgG mAb (UCH-M1; Santa Cruz Biotechnology) and visualized using the diaminobenzidine (DAB) method, as described previously (25). Four-micrometer-thick sections were mounted on poly(L-lysine)-coated slides, dewaxed, and washed in TBS (pH 7.4) for 10 min. For optimal Ag retrieval, sections were pressure cooked in 0.01 M citrate buffer (pH 6.0) for 90 s. Endogenous peroxidase activity was blocked using a 3% hydrogen peroxide solution in methanol for 10 min. Following two washes in PBS with 1% saponin, the blocking reaction was performed, as reported previously (26). Sections were incubated with primary Ab solution for 2 h at room temperature using a 1/50 dilution of the Ab. Negative control slides were incubated with rabbit IgG (R&D Systems). Secondary biotinylated anti-Ig Ab (R&D Systems) was added, and the mixture was incubated for 30 min at room temperature. Following washing, the sections were incubated with streptavidin conjugated to HRP (Amersham) and then rinsed with deionized water. DAB substrate solution was added, and the mixture was incubated for 10 min. A brown color reaction represented a positive result.
ELISA
CD14-positive cells (1 x 106 cells/ml) in BALF were incubated with 513DDR1 Ab (16, 18) or 50 µg/ml type I collagen (Sigma-Aldrich) and cultured for 24 h. Following culture, the supernatants were collected, and the concentrations of MCP-1, IL-8, MIP-1, MMP-2, MMP-9, tissue inhibitor of metalloproteinases (TIMP)-1, and TIMP-2 were measured using ELISA kits (R&D Systems), according to the manufacturer’s protocols. To evaluate the effect of 1 integrin, another collagen receptor, we used 1 integrin neutralizing mAb (DE9, 10 µg/ml; Upstate Biotechnology), as previously described (16, 18).
To evaluate whether chemokine and cytokine productions induced by DDR1 activation are dependent on p38 MAPK, we pretreated CD14-positive BALF cells with 10 µM SB203580 (Biochem-Novabiochem) for 30 min, followed by stimulation with collagen or DDR1 agonistic Ab.
We also measured MCP-1, IL-8, MIP-1, and GM-CSF concentrations in the BALF from each group.
Cytokine concentrations were determined by linear regression from a standard curve using GraphPad software (Flow Laboratories), as described previously (27).
Gelatin zymography
MMP-2 and MMP-9 activities were analyzed using gelatin zymography. Precast 7.5% polyacrylamide minigels containing 0.3% SDS and gelatin (1 mg/ml) were obtained from the YAGAI Research Center. Fifteen microliters of undiluted culture supernatant was mixed with an equal volume of 50 mM Tris-HCl buffer (pH 6.8) containing 2% SDS, 10% glycerol, and 0.01% bromphenol blue. Samples were loaded and electrophoresed at 10 mA for 20 min, followed by electrophoresis at 20 mA for 80 min, until the dye front reached the bottom of the gel. As a positive control, partially activated MMP-9 and MMP-2 samples were also loaded on the gels. Following electrophoresis, the gels were agitated in 2.5% (v/v) Triton X-100 for 1 h to remove the SDS, followed by washing in 50 mM Tris-HCl buffer (pH 7.5) containing 200 mM NaCl for 1 h to restore the enzymatic activity. The gels were incubated for 24 h at 37°C in 50 mM Tris-HCl buffer (pH 7.5) containing 200 mM NaCl, 5 mM CaCl2, 0.02% (w/v) Brij-35, and 0.01% NaN3 to allow proteolysis of the gelatin substrate. Finally, the gels were stained for 1 h with 0.1% Coomassie brilliant blue G-25 in 30% methanol and 10% acetic acid, followed by destaining for 3 h in 30% methanol and 5% acetic acid. Gelatinolytic activity was identified as clear bands against the blue background.
Western blot analysis
To detect DDR1 isoforms, 1 x 107 CD14-positive BALF cells, which were selected using magnetic beads (Miltenyi Biotec), were lysed on ice for 20 min in 1 ml of lysis buffer containing 50 mM HEPES, 150 mM NaCl, 1% Triton X-100, 10% glycerol, and a mixture of protease inhibitors (Roche). The lysates were centrifuged, and 20 µl of the supernatant was collected. Subsequently, 20 µl of double-strength sample buffer (20% glycerol, 6% SDS, and 10% 2-ME) was added to the supernatants. The samples were boiled for 10 min. Proteins were analyzed on 8% polyacrylamide gels by SDS-PAGE and transferred electrophoretically to nitrocellulose membranes at 150 mA for 1 h using a semidry system. The membranes were incubated with rabbit IgGs that specifically recognize DDR1a (16), DDR1b (18), both forms of DDR1 (Santa Cruz), or anti-human actin mouse IgG mAb (Santa Cruz Biotechnology), followed by sheep anti-rabbit or mouse IgG coupled with HRP (Amersham). Peroxidase activity was visualized by the Enhanced Chemiluminescence Detection System (Amersham). The intensities of DDR1 isoforms and actin were analyzed using the NIH Image Program (National Institutes of Health), and then the relative amount of each DDR1 isoform (DDR1 amount ratio) in each patient was calculated.
DDR1 autophosphorylation was analyzed, as previously described (16, 18, 19). First, we evaluated whether DDR1 in fresh CD14-positive BALF cells was activated by lysing 1 x 107 freshly isolated CD14-positive BALF cells, as described above. Subsequently, DDR1 in cell lysates was immunoprecipitated using anti-DDR1 Ab (C-20; Santa Cruz) and recombinant protein G-agarose (Invitrogen Life Technologies), as previously reported (16, 18, 19), and tyrosine phosphorylation of DDR1 was analyzed by Western blotting using a mouse monoclonal anti-phosphotyrosine IgG (4G10; Upstate Biotechnology) and a sheep anti-mouse IgG coupled with HRP (Amersham). Peroxidase activity was visualized by the Enhanced Chemiluminescence Detection System (Amersham).
In addition, to evaluate whether DDR1 activation by collagen or DDR1 agonistic Ab induces autophosphorylation of DDR1 and DDR1 signal transduction, 1 x 107 CD14-positive BALF cells were plated on dishes, serum starved in RPMI 1640 containing 1% FCS for 10 h, and subsequently activated with 50 µg/ml type I collagen (Sigma-Aldrich) or DDR1 agonistic Ab (513DDR1 Ab) (16, 18) and then cultured. Cell lysates were prepared, and DDR1 was immunoprecipitated and analyzed, as described above.
To detect phosphorylation of p38 MAPK, CD14-positive cells were starved, as described above, and then activated with 50 µg/ml type I collagen (Sigma-Aldrich). Twenty microliters of cell lysate was directly mixed with 20 µl of sample buffer and analyzed. Phosphorylation of p38 MAPK was analyzed by Western blotting using rabbit polyclonal anti-phosphorylated p38 Ab or rabbit polyclonal anti-p38 Ab (Cell Signaling Technology) and sheep anti-rabbit IgG HRP (Amersham). Peroxidase activity was visualized by the Enhanced Chemiluminescence Detection System (Amersham).
Statistical analysis
We used the Bonferroni-Dunn test with one-way factorial ANOVA (ANOVA). A p value below 0.05 was considered significant. Values were presented as the mean ± SD, unless stated otherwise.
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Results |
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Table I shows the clinical features of the patients investigated in this study. There was no significant difference in terms of the male/female ratio, mean age, or percentage of smokers in the three groups. The PaO2 value was significantly lower in IPF patients and COPD patients than in healthy volunteers (p < 0.05, analyzed by the Bonferroni-Dunn test with one-way factorial ANOVA). Vital capacity (VC) and DLCO% were significantly lower in IPF patients than in other groups (p < 0.01). Forced expiratory volume in 1 s (FEV1) and FEV1/VC were significantly lower in COPD patients than in other groups (FEV1, p < 0.05; FEV1/VC, p < 0.01). BALF MCP-1, IL-8, MIP-1, and GM-CSF levels in IPF patients were significantly higher than in COPD patients or healthy volunteers (p < 0.01). As shown in Fig. 1, the CD14-positive cells expressed DDR1. The percentage of DDR1-positive cells was significantly higher in IPF patients than in COPD patients or healthy volunteers. Almost all DDR1-positive cells were also CD14 positive. There was no significant difference among the three groups in the percentage of 1 integrin (another collagen receptor)-positive cells, CD14-positive cells, and 1 integrin-double-positive and CD14-positive cells in the total BALF cells (Table I).
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In all three groups, the majority of CD14-positive cells were identified as macrophages by May-Giemsa staining (IPF patients: macrophages = 95.7 ± 5.4%, neutrophils = 4.6 ± 1.2%; COPD patients: macrophages = 95.8 ± 4.5%, neutrophils = 4.5 ± 1.3%; healthy volunteers: macrophages = 96.8 ± 4.1%, neutrophils = 3.9 ± 1.1%). As shown in Fig. 1, A and B, CD14-positive cells in all three groups express DDR1; however, there was a higher frequency of CD14-positive cells in the BALF from IPF patients (Fig. 1B). The total amount of DDR1 protein was also higher per 1 x 107 cells in IPF patients (Fig. 1C), and the proportion of cells expressing the DDR1b isoform was also much higher in the BALF from these patients (Fig. 1D). This experiment was repeated using samples from five different patients, and similar results were observed.
DDR1 autophosphorylation and recruitment of Shc in freshly isolated CD14-positive BALF cells of IPF patients
To evaluate whether DDR1 activation occurs in vivo, we collected freshly isolated BALF CD14-positive cells and examined the phosphorylation levels of DDR1. As shown in Fig. 2, autophosphorylation and recruitment of Shc, the adaptor protein of DDR1b, were observed only in IPF patients and not in the other two groups. This experiment was repeated using samples from five different patients, and similar results were obtained.
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As shown in Fig. 3, infiltrating inflammatory cells in the IPF lesion stained strongly with DDR1 Ab (C-20; Fig. 3A). Bronchial and alveolar epithelial cells were moderately positive for DDR1. Infiltrating inflammatory cells were also stained positive for CD14 (Fig. 3C). Samples from three different patients showed similar results.
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As shown in Fig. 4A, stimulation with collagen or DDR1 agonistic Ab induced MCP-1, IL-8, and MIP-1 production from IPF patient’s CD14-positive BALF cells. Chemokine production was not inhibited by blocking another collagen receptor, 1 integrin. In contrast, DDR1 stimulation did not induce nor up-regulate chemokine production from CD14-positive BALF cells from either the control or COPD patients. In preliminary experiments, we cultured CD14-positive cells for various time points (4, 8, 12, 24, 36, and 48 h) and concluded that culture for 24 h is the optimal because the chemokine levels peaked at 24 h and remained elevated in IPF patients (data not shown) and, in the other groups, collagen or DDR1 agonistic Ab stimulation did not up-regulate chemokine production. Furthermore, to evaluate the effect of microbeads, we cultured the isolated CD14-positive cells and cultured freshly obtained BALF cells of each group for various time points (4, 8, 12, 24, 36, and 48 h) without any stimulation. The chemokine concentrations between isolated CD14-positive cells and freshly isolated BALF cells in the culture supernatants were compared. There was no significant difference in the chemokine concentrations between CD14-positive cell supernatants and fresh BALF cell supernatants (data not shown). Taken together, we believe that the effect of microbeads was negligible and, therefore, we concluded that up-regulated chemokine production in IPF patients was induced by DDR1 activation and not by the effect of microbeads used for the CD14-positive selection. DDR1 activation alone also up-regulated LPS-induced chemokine production from CD14-positive cells from IPF patients (Fig. 4B). Our previous study showed that DDR1b, but not DDR1a, could induce or up-regulate chemokine production from human monocyte-derived macrophages (16). In this study, CD14-positive BALF cells from COPD patients and healthy volunteers did not express DDR1b. Thus, the results of this study support the previous reports (16, 19).
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As shown in Fig. 5A, collagen or DDR1 agonistic Ab stimulation also induced MMP-9 production from IPF patient’s CD14-positive BALF cells. In preliminary experiments, we cultured CD14-positive cells for various time points (4, 8, 12, 24, 36, and 48 h) and decided that culture for 24 h is the optimal time point because the production level of MMP-9 peaked at 24 h and decreased to basal level at 48 h, while the levels of MMP-2 and TIMPs did not change (data not shown). Active MMP-9 was produced from CD14-positive BALF cells (Fig. 5B). This active MMP-9 production was not inhibited by adding an Ab that blocks 1 integrin, another collagen receptor (10 µg/ml; Upstate Biotechnology). Neither collagen nor DDR1 agonistic Ab induced MMP-2, TIMP-1, or TIMP-2 production from CD14-positive BALF cells obtained from COPD patients or healthy volunteers (data not shown).
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Based on our previous results (16, 18, 19), we predicted that DDR1b signal transduction might occur in the alveolar macrophages of IPF patient, and chose 60 min as the stimulation time. As shown in Fig. 6A, collagen stimulation induced DDR1 autophosphorylation with recruitment of Shc in IPF patients, and this was not observed in the other two groups. As shown in Fig. 6B, collagen stimulation induced p38 MAPK phosphorylation in IPF patient cells, but not in the other two groups. The results of DDR1 agonistic Ab stimulation were the same as those obtained with collagen (data not shown). These experiments were repeated using samples from five different patients, and similar results were obtained.
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To evaluate whether the chemokine and MMP-9 production induced by DDR1 activation is dependent on p38 MAPK, we pretreated CD14-positive BALF cells with a p38 MAPK inhibitor (SB203580) for 30 min and then stimulated the cells with collagen or DDR1 agonistic Ab. As shown in Fig. 7, the p38 MAPK inhibitor significantly attenuated chemokine and MMP-9 production induced by DDR1 activation. We examined the viability of the cells exposed to SB203580 using trypan blue staining and confirmed that SB203580 did not affect the viability of the cells (data not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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In this study, DDR1 activation of alveolar macrophages of IPF patients induced the secretion of chemokines, such as MCP-1, IL-8, and MIP-1. These results are compatible with our previous results (16). MCP-1 is a major chemoattractant for monocytes in inflammation and immune responses (33). MCP-1 is detectable in the BALF of IPF patients (34) and has been suggested to be associated with the pathogenesis of IPF (35). Alveolar macrophages and epithelial cells are the main cellular source of MCP-1 production in IPF (36). IL-8 is a potent chemoattractant for neutrophils and plays a pivotal role in acute inflammation by recruiting and activating neutrophils (37). As a result, IL-8 is considered to play a prominent role in the attraction of neutrophils to the lung in IPF (38). In IPF, alveolar macrophages are considered to be the cellular sources of IL-8 (39). MIP-1 is known to regulate the trafficking and activation state of select subgroups of inflammatory cells, including lymphocytes (40). MIP-1 can modulate leukocyte adhesion to the endothelium and contributes to leukocyte recruitment into the lungs (41). In addition, it was reported that BALF MIP-1 levels were elevated in IPF (42). As described above, all chemokines investigated in this study play a pivotal role in the immunological pathogenesis of IPF. The present study also showed that DDR1 activation can dramatically synergize the LPS-induced chemokine production. Taken together, we propose that DDR1 might potentially contribute to the development of IPF in combination with other factors that induce chemokine production.
Our study also showed that DDR1 activation of CD14-positive BALF cells induced the secretion of MMP-9. MMPs are a family of zinc- and calcium-dependent endopeptidases capable of proteolytically degrading many of the components of the ECM (43), and TIMPs are the endogenous inhibitors of MMPs (44). In pulmonary diseases, MMPs are believed to be associated with wound repair of the human respiratory epithelium (45), while TIMPs are believed to be associated with irreversible pulmonary structure remodeling via myofibroblasts (46). The imbalance of these factors is thought to contribute to the development of interstitial lung diseases (47). In IPF, the alveolar macrophage is one of the cellular sources that produces MMP-9 (9) and contributes to lung remodeling in IPF (31). Furthermore, DDR1 was reported to be necessary for MMP-9 production (21). Taken together, we think that DDR1 might be associated with the lung remodeling in IPF via MMP-9 production from alveolar macrophages.
Alveolar macrophages expressed higher levels of DDR1 in IPF patients than in COPD patients or healthy volunteers. The predominant isoform in IPF was DDR1b, while DDR1a was predominant in the other two groups. In our previous reports, we found that GM-CSF could induce DDR1 expression on monocytes and increase the amount of DDR1b (16, 19). In the present study, the GM-CSF concentration in BALF was significantly higher in IPF patients than in other groups. Therefore, we think that the high GM-CSF concentration in BALF might be one of the reasons for the difference in total DDR1 levels and in the DDR1 isoform expression pattern between IPF patients and the other groups. Furthermore, alveolar macrophages of IPF patients are considered to differ functionally and metabolically from alveolar macrophages obtained from normal subjects because previous studies reported the spontaneous release of chemotactic factors for recruiting inflammatory cells (48, 49). The difference in the DDR1 isoform expression pattern in alveolar macrophages might be associated with the functional difference in macrophages of IPF patients and healthy individuals.
DDR1 activation can induce phosphorylation of p38 MAPK via a unique signaling pathway (16, 18, 19). In addition, monocyte-derived macrophages secrete MCP-1, IL-8, and MIP-1 following DDR1 activation in a p38 MAPK-dependent manner (16). The p38 MAPK is activated in the lung tissue of IPF patients (50), and also in a murine IPF model (51). In addition, the inhibition of p38 MAPK can ameliorate murine bleomycin-induced pulmonary fibrosis (52). Thus, p38 MAPK involvement in the pathogenesis of IPF is known; however, there is little information regarding the signaling pathway upstream of p38 MAPK. In our report, DDR1-induced chemokine and MMP-9 production in alveolar macrophages was dependent on the p38 MAPK pathway. Therefore, we suggest that DDR1 might be one of the upstream signaling molecules of p38MAPK in IPF.
The cell surface receptor, 1 integrin, is another well-known collagen receptor, and ligation of 1 integrin can induce the expression of proinflammatory cytokines in monocytes (53). However, the expression of 1 integrin is low or undetectable on monocyte-derived macrophages (54, 55), suggesting the presence of collagen receptors other than 1 integrins that may promote cytokine production in macrophages. Indeed, the present study showed a significant difference in the DDR1 expression level between IPF patients and healthy volunteers or COPD patients, while there was no significant difference between the percentage of 1 integrin-positive alveolar macrophages. Taken together, we think that DDR1, not 1 integrin, is the collagen receptor on alveolar macrophages that contributes to the pathogenesis of IPF.
In conclusion, we showed functional DDR1b expression on alveolar macrophages of IPF patients and proposed a possible association of DDR1 in the pathogenesis of IPF. To date, there have been many reports investigating the inflammatory mediators or molecular interaction between inflammatory mediators and inflammatory cells in IPF; however, there is little information regarding the molecular interaction between the microenvironment, such as ECM, and inflammatory cells. Therefore, further studies addressing this point are necessary to clarify the pathogenesis of IPF.
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Acknowledgments |
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Disclosures |
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Footnotes |
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1 This study was supported by a grant-in-aid for scientific research (16790447) from Japan Society for the Promotion of Science, a grant from the Sumitomo Foundation (040010), a grant from Nagao Memorial Fund, and a grant from Uehara Memorial Foundation. 
2 Address correspondence and reprint requests to Dr. Wataru Matsuyama, Division of Respiratory Medicine, Respiratory and Stress Care Center, Kagoshima University Hospital, Sakuragaoka 8-35-1, Kagoshima 890-8520, Japan. E-mail address: vega{at}xa2.so-net.ne.jp
3 Abbreviations used in this paper: IPF, idiopathic pulmonary fibrosis; BALF, bronchoalveolar lavage fluid; COPD, chronic obstructive lung disease; DAB, diaminobenzidine; DDR1, discoidin domain receptor 1; ECM, extracellular matrix; FEV1, forced expiratory volume in 1 s; MMP, matrix metalloproteinase; TIMP, tissue inhibitor of metalloproteinases; VC, vital capacity.
Received for publication November 1, 2004. Accepted for publication March 2, 2005.
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) promotes migration of leukocytes in three-dimensional collagen lattices. FASEB J. 15: 2724-2726.
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 C. Agostini and C. Gurrieri Chemokine/Cytokine cocktail in idiopathic pulmonary fibrosis. Proceedings of the ATS, January 1, 2006; 3(4): 357 - 363. [Abstract] [Full Text] [PDF]
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W. Matsuyama, H. Mitsuyama, M. Watanabe, Y. Shirahama, I. Higashimoto, M. Osame, and K. Arimura Involvement of Discoidin Domain Receptor 1 in the Deterioration of Pulmonary Sarcoidosis Am. J. Respir. Cell Mol. Biol., December 1, 2005; 33(6): 565 - 573. [Abstract] [Full Text] [PDF]
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