Abstract
Tobacco smoke is believed to cause small airway disease and then chronic obstructive pulmonary disease (COPD), but the molecular mechanisms by which small airway obstruction occurs remain unknown. To study the gene expression levels of transforming growth factor (TGF)- β 1, a potent fibrogenic factor, in small airway epithelium from smokers and patients with COPD, we harvested highly pure samples of epithelial cells from small airways under direct vision by using an ultrathin bronchofiberscope BF-2.7T (outer diameter 2.7 mm with a biopsy channel of 0.8 mm in diameter). The expression levels of TGF- β 1 were evaluated by reverse transcription-polymerase chain reaction (RT-PCR). The mRNA levels of TGF- β 1 corrected by β -actin transcripts were significantly higher in the smoking group and patients with COPD than those in nonsmokers (p < 0.01). Furthermore, among smokers and patients with COPD, TGF- β 1 mRNA levels correlated positively with the extent of smoking history (pack-years) and the degree of small airway obstruction as assessed by measurements of flow–volume curves. Immunocytochemistry of the cells demonstrated more intense stainings for TGF- β 1 in samples from smokers and patients with COPD than from nonsmokers. Spontaneously released immunoreactive TGF- β 1 levels from cultured epithelial cells were more elevated in subjects with a history of smoking and patients with COPD than in nonsmokers. Our study showed a close link between smoking and expression of TGF- β 1 in small airways. Our results also suggested that small airway epithelial cells might be involved in obstructive changes found in smokers and patients with COPD.
Tobacco smoke has been implicated as one of the most important factors that can cause small airway disease followed by chronic obstructive pulmonary disease (COPD) (1, 2). Animal models of tobacco exposure showed inflammatory responses along the airways as well as lung parenchymal structures (3– 5), but the molecular mechanisms by which small airway obstruction occurs remain unknown. Recent studies suggest some fibrogenic growth factors may be involved in the remodeling processes of the small airways (6, 7). One of the most potent and extensively studied growth factors is transforming growth factor (TGF)-β1, which induces fibroblast proliferation, increased production of collagen and other extracellular matrix proteins, and decreased collagen degradation (8). This growth factor is also chemotactic for macrophages (9) and mast cells (10). TGF-β1 has an inhibitory effect on growth of epithelial cells including bronchial epithelial cells (11, 12). TGF-β1 is usually released as inactive forms, which are activated by cleavage of SH-bonds with proteolytic enzymes. Airway epithelial cells constitutively express these polypeptides in both active and inactive forms, and thereby can function as autocrine growth inhibiting factors for themselves (13, 14). TGF-β1 mRNA and proteins are localized within peripheral bronchial epithelial cells in normal lungs (15). de Boer and coworkers (16) recently demonstrated that TGF-β1 mRNA and proteins were increased in peripheral lung tissues from smokers with and without COPD by immunostaining and in situ hybridization techniques. Peripheral airway epithelial cells are, therefore, important possible sources of this growth factor and may play a role in airway mucosal repair and increased collagen deposition along the airway walls.
We harvested living epithelial cells from small airway mucosa under direct vision by using an ultrathin bronchofiberscope BF-2.7T (outer diameter 2.7 mm with a biopsy channel 0.8 mm in diameter) (17, 18). Although we found that expression of interleukin-8 (IL-8) and ICAM-1 was significantly higher in small airway epithelium from tobacco smokers than from nonsmokers (19), there was no correlation between these inflammatory markers and airway obstruction in small airways as assessed by V˙25.
In the present study, we evaluated the mRNA levels of TGF-β1 in small airway epithelium from nonsmokers, smokers, and patients with COPD by the reverse transcription and polymerase chain reaction (RT-PCR) technique, and correlated the magnitude of the expression of this fibrogenic factor to the degree of small airway obstruction.
Thirty-three Japanese healthy volunteers and 17 patients with COPD were included in the study. Among healthy volunteers, 18 people (14 males and 4 females, mean age 58.0 yr) were current smokers and 15 people were never smokers (12 males and 3 females, mean age 53.3 yr). All the subjects were free from respiratory diseases, with normal chest X-ray films and with no respiratory symptoms for at least 3 mo before this study. Spirometry was performed in all subjects with no abnormal results in percentage FVC and FEV1 among healthy people. Maximal flow rates at 50% and 25% lung volumes (V˙50 and V˙25, respectively) were also evaluated and expressed as percent predicted values (20). The diagnosis of COPD basically depended on the reported criteria (16) including decreased percentage FEV1 (< 75%) and no response to inhalation of bronchodilators. The data for V˙25 were significantly greater in never smokers than in smokers and patients with COPD (p < 0.01, ANOVA). The clinical data of these subjects are summarized in Table 1. The study was planned according to the ethical guidelines following the declaration of Helsinki and given institutional approval and an informed consent was obtained from each subject.
| Age (yr) | Sex | Smoking History | Pulmonary Function Tests | |||||||||||||
|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
| FVC (%) | FEV1 (%) | V˙ 50 (%) | V˙ 25 (%) | V˙ 50/V˙ 25 | ||||||||||||
| Nonsmokers (n = 15) | 53.3 ± 1.68* | M12:F3 | — | 91.4 ± 2.10 | 81.0 ± 4.21 | 85.4 ± 2.82 | 66.5 ± 3.80 | 1.89 ± 0.10 | ||||||||
| Smokers (n = 18) | 56.1 ± 1.90 | M14:F4 | 42.1 ± 4.50 | 90.4 ± 4.16 | 82.2 ± 3.01 | 85.1 ± 2.90 | 48.0† ± 3.60 | 2.99† ± 0.28 | ||||||||
| COPD (n = 17) | 58.0 ± 2.10 | M14:F3 | 44.7 ± 3.52 | 93.5 ± 3.54 | 65.2†,‡ ± 2.91 | 64.2†,‡ ± 4.30 | 35.9† ± 2.60 | 3.23† ± 0.28 | ||||||||
The subjects underwent a bronchofiberscopic examination with a BF-XT20 fiberscope (Olympus, Tokyo, Japan) in a standard fashion (17, 18). Under fluorographic guidance an ultrathin fiberscope (BF-2.7T) was inserted through a 2.8-mm-diameter biopsy channel. A newly modified BC-0.7T brush was then inserted to collect cells by brushing the airway mucosal surfaces several times. Brushing of the mucosa was routinely performed at three or four different ninth or tenth lower lobe bronchioles. The cells were immediately collected by vortexing the brush in RPMI 1640 medium supplemented with 10% fetal calf serum (FCS, heat inactivated; GIBCO, Grand Island, NY). The cells were centrifuged for 5 min at 1000 rpm. The recovered cells were washed twice in Hanks' balanced salt solution without calcium and magnesium (HBSS; GIBCO). The number of the cells was counted by a standard hemocytometer and the cell viability was assessed by the trypan blue dye exclusion technique (18).
The cytospin preparations from harvested cells were obtained by a cytocentrifuge, and were routinely stained by Diff-Quick stain (modified Wright–Giemsa stain; Midorijuji, Kobe, Japan). The cytospin preparations were also stained by periodic acid-Schiff (PAS) stain for the detection of secretory granules. For detection of keratin in the cells, the specimens were stained with antikeratin (KL-1; Immunotech, Marseille, Cedex, France), or with control IgG1 monoclonal antibodies using the avidin-biotin complex method (21, 22). The differential counts of the harvested cells basically depended on the Diff-Quik and PAS stainings and were divided into four categories: ciliated, secretory, and nonciliated epithelial cells as well as other inflammatory cells (23).
To assess the TGF-β1 mRNA levels in human small airway epithelial cells, a semiquantitative assay utilizing RT-PCR was performed as previously reported (18, 24). We used the epithelial cell samples for RT-PCR only when the samples contained less than 5% nonepithelial cells as evaluated by Diff-Quik and keratin staining. Total RNA was isolated by the guanidinium thiocyanate–phenol–chloroform extraction method as described by Chomczynski and Sacchi (25). Briefly, after cell counting and assessment of cell viability, the cells (5.0 × 105 viable cells) were lysed in solution D (4 M guanidinium thiocyanate, 25 mM sodium citrate, pH 7; 0.5% sarcosyl, 0.1 M 2-mercaptoethanol) and RNA was extracted from the solution by chloroform extraction. After that, the isopropanol precipitate RNA was washed twice with 70% ethanol, dried, and resuspended in diethylpyrocarbonate-treated water. Extracted RNA was reverse transcribed to cDNA by using a Takara RNA-PCR kit according to the manufacturer's recommendations. Briefly, total RNA, random hexadeoxynucleotides as primer, and avian myeloblastosis virus reverse transcriptase were used for cDNA synthesis. The following specific primer pairs were used for PCR amplification:
TGF-β1 (5′primer) 5′-GCCCTGGACACCAACTATTGCT-3′
(3′primer) 5′-AGGCTCCAAATGTAGGGGCAGG-3′
β-actin (5′primer) 5′-ATCTGGCACCACACCTTCTACAATGAGCTGCG-3′
(3′primer) 5′-CGTCATACTCCTGCTTGCTGATCCACATCTGC-3′
(Clontech, Palo Alto, CA).
The reaction mixture contained 10 mM Tris–HCl (pH 8.3 at 25° C), 50 mM KCl, 1.5 mM MgCl2, 1 mg/ml gelatin, 0.4 μM of each primer, 0.25 M diethyl-p-nitrophenyl monothiophosphate (dNTP), 1.0 μg cDNA, and 1 U of Taq polymerase (Perkin-Elmer-Cetus, Norwalk, CT) in 25 μl. Amplification was performed for allotted cycles of denaturing (94° C, 2 min), annealing (60° C, 30 s), and extension (72° C, 1.5 min) using a thermal cycler (Progene; Techne, Cambridge). The PCR cycle was determined by preliminary experiments showing a linear relationship between PCR cycles and intensity of signals on ethidium bromide-stained agarose gels. For semiquantitative evaluation of TGF-β1 and β-actin mRNAs, 30 and 25 cycles were chosen, respectively. PCR product was run on a 1.0% agarose gel, and the intensity of ethidium bromide fluorescence was evaluated by NIH Image version 1.61.
It was possible that the results of RT-PCR described above might be influenced by an artifact during the amplification processes. To improve confidence in the results, we reevaluated the mRNA levels of β-actin and TGF-β1 by doing the amplification processes in the same tube. In these experiments, PCR amplification processes were set to 25 cycles.
In some experiments, where enough epithelial cells were obtained, the cells were plated onto collagen-coated 48-well flat-bottom tissue culture plates (Koken, Tokyo, Japan) at a density of 5 × 104 cells/well with hormonally defined SAGB medium (Clonetics; SankoJunyaku Co., Ltd., Tokyo, Japan). Morphological changes during culture were studied by phase-contrast microscopy showing polygonal, nonciliated cells with a tight connection to each other. Confluent monolayers of epithelial cells were stained with antikeratin (KL-1; Immunotech), antivimentin (DAKO-Vimentin; DAKOPatts, Glostrup, Denmark), or with control IgG1 monoclonal antibodies using the avidin-biotin complex method (21, 22) to show that the cells were of epithelial cell origin.
On confluency, the epithelial cell-conditioned media were harvested after different time periods. Immunoreactive TGF-β1 was measured by specific ELISA (R&D Systems, Inc., Minneapolis, MN) and was expressed in pg/106 cells/24 h. For measurement of total amounts of TGF-β1, the samples were acidified by the addition of 1 N HCl followed by neutralization with NaOH as recommended by the manufacturer.
The cell samples were cytocentrifuged onto glass slides by Cytospin 2 (Shandon Southern Products, Cheshire, England) and fixed with 4% paraformaldehyde. Immunocytochemistry was performed using avidin-biotin complex peroxidase (ABO-PO) as described previously in detail (26, 27). Briefly, the fixed cells were preincubated with 10% normal goat serum (Nichirei Corp., Tokyo, Japan) for 15 min to prevent nonspecific binding. The cells were then incubated with an anti-TGF-β1-specific antibody (1:100 diluted, 10 μg/ml; R&D Systems) for 30 min at room temperature. After rinsing in phosphate-buffered saline (PBS), the slides were incubated with biotin-conjugated goat anti-rabbit immunoglobulin G antibody (Nichirei) and ABC-PO solution for 30 min. After rinsing in PBS, the reaction products were visualized using diaminobenzidine (Sigma). The staining intensity was graded and expressed as 0 = absence of staining, 1 = moderate staining, 2 = intense staining, and 3 = very intense staining, as reported by de Boer and coworkers (16).
The results were analyzed by nonparametric equivalents of analysis of variance (ANOVA) for multiple comparison as reported (18). Spearman's rank correlation test was used for correlation analysis between the two data.
The total cell number and cell differentials evaluated by Diff-Quick, PAS and keratin stainings were shown in Table 2. The total cell number, viability and cell differential counts were not statistically different among the non-smokers, smokers and patients with COPD. The number of the recovered cells ranged from 1.20 × 106 to 2.30 × 106 with the mean of 1.78 × 106. The cell viability ranged from 62.5% to 79.5% with the mean of 68.0%. Approximately 70% of the viable cells were non-ciliated round cells which were positive to keratin staining as reported previously (18). As shown in Table 2, the major contaminating cells were neutrophils, but the percent of the non-epithelial cells were less than 5% in all cases.
| Cell number (×106 ) | Viability (%) | Keratin (+) Cells (%) | Cell Differentials* | |||||||||||||||
|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
| Ciliated Cells (%) | Secretory Cells (%) | Nonciliated Cells (%) | Neutrophils (%) | Eosinophils (%) | Macrophages (%) | |||||||||||||
| Nonsmokers (n = 15) | 1.56 ± 0.65† | 67.3 ± 5.00 | 95.0 ± 7.35 | 21.9 ± 5.95 | 3.95 ± 4.05 | 71.0 ± 4.25 | 2.00 ± 1.00 | 0.25 ± 0.03 | 0.90 ± 0.55 | |||||||||
| Smokers (n = 13) | 1.85 ± 0.55 | 71.8 ± 2.75 | 97.0 ± 3.01 | 20.0 ± 3.35 | 5.99 ± 3.20 | 69.5 ± 2.70 | 2.75 ± 0.95 | 0.15 ± 0.33 | 1.61 ± 0.70 | |||||||||
| COPD (n = 12) | 1.80 ± 0.95 | 69.0 ± 5.30 | 95.0 ± 1.90 | 21.6 ± 4.00 | 5.35 ± 2.09 | 69.2 ± 3.55 | 2.65 ± 1.90 | 0.23 ± 0.05 | 1.50 ± 0.85 | |||||||||
The signals for TGF-β1 and β-actin were detected in all cases, and the relative intensity of TGF-β1 mRNA/β-actin was statistically higher in smokers and patients with COPD than in nonsmokers as shown in Figure 1a and 1b. The TGF-β1 mRNA levels in patients with COPD were significantly higher than those of healthy smokers.
Fig. 1. TGF-β1 mRNA levels in the small airway epithelial cells obtained from nonsmokers, tobacco smokers, and patients with COPD as evaluated by RT-PCR analysis. (a) A representative result of RT-PCR gel showing that the magnitude of TGF-β1 signals from smokers and those with COPD was stronger than from nonsmokers (n = 3 in each group). P, positive control; M, molecular marker. (b) Semiquantitative evaluation of the TGF-β1 mRNA levels corrected by β-actin transcripts indicating that the signals from smokers (n = 13) and patients with COPD (n = 12) were statistically stronger than those from nonsmokers (n = 10). Patients with COPD showed a significantly stronger signal as compared with smokers without airway obstruction. *p < 0.01, ANOVA. (c) Semiquantitative evaluation of the TGF-β1 mRNA levels corrected by β-actin transcripts among samples amplified in the same tubes. The signals from smokers (n = 11) and patients with COPD (n = 8) were statistically stronger than from nonsmokers (n = 7). Patients with COPD showed a significantly stronger signal as compared with smokers without airway obstruction. *p < 0.01, ANOVA. (d) RT-PCR for TGF-β1 mRNA levels using attached cells. The epithelial cell samples were plated onto collagen-coated culture dishes to allow attachment. The attached cells were more than 98.5% positive to keratin staining. Semiquantitative evaluation of the TGF-β1 mRNA levels corrected by β-actin transcripts among samples amplified in the same tubes. The signals from smokers (n = 5) and patients with COPD (n = 5) were statistically stronger than from nonsmokers (n = 5). Patients with COPD showed a significantly stronger signal as compared to smokers without airway obstruction. *p < 0.05, ANOVA.
We reevaluated the mRNA levels of TGF-β1 and β-actin in the same tube when a large enough amount of cDNA was obtained. Among 7 nonsmokers, 11 smokers, and 8 patients with COPD, the levels of TGF-β1 corrected by β-actin transcripts were again statistically higher in smokers and in patients with COPD than in nonsmokers (Figure 1c).
To exclude the possibility that the above observations were caused by the contamination of a few nonepithelial cells such as neutrophils, we incubated the cells on collagen-coated tissue culture plates for 90 min to allow the epithelial cells to attach, and then the plates were rinsed twice and RNA was extracted (n = 5 in each group among never smokers, smokers, and patients with COPD). By this technique, the keratin-positive cells were always more than 98.5% in all the samples as assessed by immunostain on collagen-coated LabTec chamber slides. The results again elucidated that the signals for TGF-β1 mRNA were significantly increased in smokers and in patients with COPD than in nonsmokers (Figure 1d).
Among current smokers without airway obstruction, TGF-β1 mRNA levels correlated positively with the extent of smoking history when the signals were normalized by β-actin transcripts (r = 0.620, p < 0.001) (Figure 2a). The magnitude of TGF-β1 expression in patients with COPD also showed a significant correlation with smoking history as shown in Figure 2b (r = 0.653, p < 0.001). Such was also the case when the data from PCR in the same tubes were evaluated (data not shown).
Fig. 2. Correlation between the magnitude of TGF-β1 mRNA/β-actin transcripts and the burdens of smoking (pack-years). There was a significant positive correlation by Spearman's rank correlation analysis in (a) healthy smokers (r = 0.620, p < 0.001) and (b) patients with COPD (r = 0.653, p < 0.001).
[More] [Minimize]The mRNA levels of TGF-β1 significantly correlated with percentage V˙25 and V˙50/V˙25 in smokers (Figure 3a and 3c), but there was no relationship to percentage V˙50 (Figure 3b), FVC, or FEV1 (data not shown). Levels in patients with COPD showed significant correlation with percentage V˙50 as well as V˙25, but not with V˙50/V˙25 (Figure 4), FVC, or FEV1 (data not shown). Such was also the case when the data from PCR in the same tubes were evaluated (data not shown).
Fig. 3. Correlation between the magnitude of TGF-β1 mRNA/β-actin transcripts and pulmonary function test results among healthy smokers. There was a significant correlation between TGF-β1 expression and (a) V˙ 25 and (c) V˙ 50/V˙ 25, but not (b) V˙ 50 by Spearman's rank correlation analysis.
[More] [Minimize]
Fig. 4. Correlation between the magnitudes of TGF-β1 mRNA/β-actin transcripts and pulmonary function test results among patients with COPD. There was a significant correlation between TGF-β1 expression and (a) V˙ 25 and (b) V˙ 50, but not with V˙ 50/V˙ 25 by Spearman's rank correlation analysis.
[More] [Minimize]The recovered epithelial cells were subject to TGF-β staining. As shown in Figure 5, airway epithelial cells from smokers and patients with COPD showed more intense staining than from nonsmokers.
Fig. 5. Immunocytochemistry for TGF-β1 in small airway epithelial cells. (a) Photomicrographs showing immunostaining of the cells from nonsmokers, smokers, and patients with COPD. (b) A semiquantitative evaluation of TGF-β1 protein levels in small airway epithelial cells. The scores from smokers (n = 8) and patients with COPD (n = 9) were statistically greater than from nonsmokers (n = 8). *p < 0.05.
[More] [Minimize]The cells from nonsmokers (n = 5), smokers (n = 5), and patients with COPD (n = 6) were cultured until confluence in SAGB medium. Immunocytochemical studies demonstrated that the cells were keratin positive, but vimentin negative, showing the cells were epithelial cells. Inactive and active forms of TGF-β1 proteins spontaneously released by the epithelial cells were evaluated after different time periods by specific enzyme-linked immunosorbent assay (ELISA). As shown in Figure 6, there was a time-dependent accumulation of immunoreactive TGF-β1 in culture supernatants and total (inactive and active) amounts of TGF-β1 were greater in smokers and patients with COPD than in nonsmokers after 24 h. However, the levels of active form of TGF-β1 were not different among the three groups. We also studied the effects of stimulation with IL-1β (10 ng/ml) and tumor necrosis factor (TNF)-α (10 ng/ml) in some samples. Neither stimulus affected the release of TGF-β1 (data not shown).
Fig. 6. Spontaneous release of active and inactive forms of TGF-β1 proteins from small airway epithelial cells in culture. Total amounts released from small airway epithelial cells were significantly increased in smokers (n = 5) and patients with COPD (n = 6) than in nonsmokers (n = 5). TGF-β1 release in patients with COPD was greater than in smokers. The amounts of active form of TGF-β1 did not change among three groups. *p < 0.01 as compared with nonsmokers; +p < 0.01 as compared with smokers without airflow limitation, ANOVA. n.s., not significant.
[More] [Minimize]In the present studies, we found that the levels of TGF-β1 mRNA in small airway epithelium were significantly higher in smokers and patients with COPD compared with nonsmokers. Importantly, the magnitude of the TGF-β1 signals showed a positive correlation with burden of cigarette smoking. Our results further showed that TGF-β1 gene expression levels correlated with the degrees of peripheral airway obstruction as evaluated by measurements on flow–volume curves. Immunocytochemistry also showed that the TGF-β1 protein was more intensely detected in epithelial cells from smokers and patients with COPD than from nonsmokers. Spontaneous release of total TGF-β1 protein from cultured epithelial cells was also increased in tobacco smokers and patients with COPD compared with nonsmokers.
It is well known that cigarette smoking causes inflammatory responses in small airways (1, 2, 28). These changes included infiltration of inflammatory cells such as neutrophils, macrophages, and mast cells, and thickening of the airway walls with increased collagen deposition (1, 2). Such inflammatory and remodeling processes are believed to be related to the obstruction in small airways. The tobacco-related changes in the peripheral airways impose a major risk for development of COPD, especially pulmonary emphysema. Local migration of neutrophils might be induced by the direct effect of tobacco contents (29), however, additional data suggested that cigarette smoke stimulated airway epithelial cells to release chemotactic activities for neutrophils (30, 31) such as IL-8. In fact, the mRNA levels of IL-8 and one of the important adhesion molecules ICAM-1 were increased in small airway epithelial cells from smokers as compared with nonsmokers (19). However, it remained unclear whether these cells were also involved in the steps of tissue repair and remodeling, and eventual obstructive changes in the small airways. Recently, Stefano and coworkers (30) demonstrated that severity of airflow limitation is associated with severity of airway inflammation in smokers.
TGF-β has been hypothesized to be involved in airway remodeling found in chronic airway inflammatory disorders such as COPD and asthma (31, 32). de Boer and coworkers (16) studied the expression of TGF-β1 mRNA and proteins in resected lungs from smokers and patients with COPD by in situ hybridization and immunostaining techniques. They showed that semiquantitative histological scores of TGF-β1 mRNA and protein levels assessed by visual analogue scoring system were significantly increased in bronchiolar and alveolar epithelium as well as endothelium in these subjects. There were significant correlations between the scores of TGF-β1 mRNA or proteins in bronchiolar epithelium, and the number of macrophages or mast cells, suggesting a role of this growth factor in the accumulation of these inflammatory cells. Aubert and coworkers (33) studied the expression levels of TGF-β1 in lungs from patients with asthma, nonobstructive tobacco smokers, and patients with COPD by Northern blot analysis. They found no difference among the three study groups. These apparent discrepancies of these results might be due to the different techniques utilized for evaluation. Aubert's group studied the magnitude of mRNA by Northern blot analysis using lung tissues, and de Boer and coworkers (16) used in situ hybridization and immunostaining techniques. In the present studies, we obtained highly pure populations of small airway epithelial cells by an ultrathin scope from volunteers and patients with COPD who were free of lung cancer or other lung diseases necessitating surgery. Then, we compared the levels of TGF-β1 mRNA among nonsmokers, smokers without COPD, and patients with COPD. The levels of TGF-β1 mRNA were statistically increased in people who smoked compared with nonsmokers. We also studied the TGF-β1 protein expression by immunocytochemical analysis, and observed results comparable to the findings of de Boer and coworkers (16). Total amounts of spontaneously released TGF-β1 were also increased in smokers and patients with COPD, but the active forms of this peptide did not change among the three groups, possibly because its activation is mainly dependent on proteolytic enzymes in the local microenvironments.
Our present findings needed careful considerations, as there were several methodological limitations. First, RT-PCR itself was not a quantitative technique for the evaluation of certain gene expression levels to reconfirm the findings. To improve the technique, we repeatedly evaluated expression levels by amplifying β-actin and TGF-β1 genes in the same tubes. We also studied the expression of TGF-β1 protein by immunocytochemical analysis. Although it was difficult to evaluate the intensity of staining, these studies again showed increased TGF-β1 expression in peripheral bronchial epithelial cells when a visual analogue scale reported previously (16) was applied. Second, a small number of nonepithelial cells in the cell samples might have led to the misleading results. To exclude this possibility, attached cells were used for RT-PCR analysis. The cells were more than 98.5% epithelial cells and the results again showed that the levels of TGF-β1 gene expression were higher in the epithelial cells from smokers and patients with COPD than from nonsmokers. Immunocytochemistry also demonstrated that the positive cells were virtually all epithelial cells. In accordance with the findings by RT-PCR, increased release of total TGF-β1 from cultured epithelial cells was found in smokers and patients with COPD. However, the cultured epithelial cells might have been stimulated by attachment to the culture plates, or might have been changed as a result of being retrieved from microenvironments in the airways. Results of immunostaining showed that the freshly recovered epithelial cells from smokers and patients with COPD showed increased staining as compared with nonsmokers, which was in accordance with the findings from cultured epithelial cells. Other approaches such as an in situ hybridization technique will better elucidate gene expression in the specific cell types in the airways.
In conclusion, we demonstrated an increased expression of TGF-β1 mRNA in small airway epithelium from healthy smokers and patients with COPD. It was suggested that small airway epithelial cells might be involved in the processes of airway remodeling and the resultant obstructive changes in the small airways.
Supported in part by the Adult Diseases Memorial Foundation and the Manabe Medical Foundation.
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