Abstract
Chronic obstructive pulmonary disease (COPD) is characterized by infiltration of inflammatory cells, destruction of lung parenchyma, and airway wall remodeling. Hyaluronan (HA) is a component of the extracellular matrix, and low-molecular-weight (LMW) HA fragments have proinflammatory capacities. We evaluated the presence of HA in alveolar and airway walls of C57BL/6 mice that were exposed to air or cigarette smoke (CS) for 4 weeks (subacute) or 24 weeks (chronic). We measured deposition of the extracellular matrix proteins collagen and fibronectin in airway walls and determined the molecular weight of HA purified from lung tissue. In addition, we studied the expression of HA-modulating genes by RT-PCR. HA staining in alveolar walls was significantly enhanced upon chronic CS exposure, whereas HA levels in the airway walls were already significantly higher upon subacute CS exposure and remained elevated upon chronic CS exposure. This differed from the deposition of collagen and fibronectin, which are only elevated at the chronic time point. In lungs of CS-exposed mice, the molecular weight of HA clearly shifted toward more LMW HA fragments. CS exposure significantly increased the mRNA expression of the HA synthase gene Has3 in total lung tissue, whereas the expression of Has1 was decreased. These in vivo studies in an experimental model of COPD show that CS exposure leads to enhanced deposition of (mostly LMW) HA in alveolar and bronchial walls by altering the expression of HA-modulating enzymes. This may contribute to airway wall remodeling and pulmonary inflammation in COPD.
Chronic obstructive pulmonary disease (COPD) is a disease of the airways and lungs that is characterized by a progressive airflow limitation that is not fully reversible (1). The airflow limitation is associated with an abnormal inflammatory response of the lungs to noxious particles and gases. Significant extrapulmonary effects also contribute to the severity of the disease (www.goldcopd.com). The prevalence of COPD is increasing, and COPD will soon be the third leading cause of death worldwide (2, 3). The pathology of COPD includes obstruction of the small airways (bronchiolitis) and destruction of the lung parenchyma (emphysema). Patients with COPD also show pathologically distinct structural alterations of the small airways (airway wall remodeling) (4). The molecular and cellular mechanisms leading to cigarette smoke (CS)-induced inflammation, pulmonary emphysema, and airway wall remodeling have not been elucidated. Cigarette smoking is by far the most important risk factor for the development of COPD, and it has been shown that chronic exposure to CS leads to lung inflammation with increased numbers of inflammatory cells, including macrophages (5, 6), dendritic cells (DCs) (7–9), neutrophils (10, 11), and CD8+ T lymphocytes (12).
Pulmonary emphysema and airway wall remodeling in COPD are associated with damage and ineffective repair of the extracellular matrix (ECM). Hyaluronan (HA) is a component of the ECM that is present at enhanced levels in sputum of patients with COPD (13). HA exists in healthy tissues as a high-molecular-weight (HMW) nonsulfated glycosaminoglycan polymer, composed of repeating disaccharide units of glucuronic acid and N-acetylglucosamine. In the ECM, HA plays a role in water homeostasis, plasma protein distribution, and matrix structure (14). HA has additional biological functions that are reported to be dependent on its molecular size. Oligosaccharides of HA can induce angiogenesis and endothelial cell proliferation (15) and can activate DCs (16). Low-molecular-weight (LMW) HA fragments, with an average molecular mass of 250 kD, exhibit proinflammatory capacities by stimulating production of cytokines, chemokines, and matrix metalloproteinase–12 by macrophages (17, 18). In contrast, HMW HA has protective effects for osteoarthritis and rheumatoid arthritis (19). Under normal conditions, there is a continuous turnover of HA. Three isoforms of human HA synthases have been identified, all differing in their enzymatic properties and the length of the HA chain that is formed (20). HA synthase (Has)1 and Has2 synthesize HMW HA, whereas Has3 creates LMW HA fragments. HA is broken down by hyaluronidases, of which hyaluronidase (Hyal)1 and Hyal2 are the most important in somatic tissues. Hyal2 degrades HMW HA to LMW HA fragments, whereas Hyal1 cleaves LMW HA to oligosaccharides (21).
In this study, we report the effects of subacute and chronic CS exposure on alveolar and bronchial deposition of the glycosaminoglycan HA in a murine model of COPD. We measured the bronchial deposition of the ECM proteins collagen and fibronectin and compared the time courses of the CS-induced modulation of deposition of HA versus ECM proteins. We determined the molecular weight of hyaluronan in lungs of air- and CS-exposed mice. In addition, we analyzed the in vivo effect of CS on mRNA expression of HA synthases and hyaluronidases in total lung tissue and isolated lung macrophages.
Male C57BL/6 mice, 6 to 8 weeks old, were purchased from Harlan (Zeist, The Netherlands). The local Ethics Committee for animal experimentation of the faculty of Medicine and Health Sciences (Ghent, Belgium) approved all in vivo manipulations.
Mice (n = 8 per group) were exposed whole body to CS as described previously (22–24). Briefly, groups of eight mice were exposed to the tobacco smoke of five cigarettes (Reference Cigarette 2R4F without filter; University of Kentucky, Lexington, KY) four times a day with 30-minute smoke-free intervals 5 days per week for 4 weeks (subacute exposure) or 24 weeks (chronic exposure). An optimal smoke/air ratio of 1:6 was obtained. The control groups were exposed to air. Carboxyhemoglobin in serum of CS-exposed mice reached a nontoxic level of 8.3 ± 1.4%, compared with 1.0 ± 0.2% in air-exposed mice (n = 7 for both groups), which is similar to carboxyhemoglobin blood concentrations of human smokers (25).
Twenty-four hours after the last smoke exposure, mice were weighed and killed with an overdose of pentobarbital (Sanofi, Libourne, France), and a tracheal cannula was inserted. Three fractions of 300 μl followed by three fractions of 1 ml of Hanks' balanced salt solution, free of ionized calcium and magnesium but supplemented with 0.05 mM sodium EDTA, were instilled via the tracheal cannula and recovered by gentle manual aspiration. The six lavage fractions were pooled and centrifuged, and the cell pellet was washed twice and resuspended in 1 ml of Hanks' balanced salt solution. A total cell count was performed in a Bürcker chamber, and the differential cell counts (on at least 400 cells) were performed on cytocentrifuged preparations using standard morphologic criteria after May-Grünwald-Giemsa staining. Flow cytometric analysis of bronchoalveolar lavage (BAL) cells was performed to enumerate DCs and CD4+ and CD8+ T cells.
Cells were preincubated with Fc-receptor blocking antibody (anti-CD16/CD32, clone 2.4G2) to reduce nonspecific binding. Monoclonal antibodies used to identify mouse DC populations were allophycocyanine (APC)-conjugated anti-CD11c (N418) and phycoerythrin-conjugated anti–I-Ab (AF6-120.1). We discriminated between macrophages and DCs using the methodology described by Vermaelen and colleagues (26). After gating on the CD11c-bright population, two peaks of autofluorescence can be distinguished. Macrophages are identified as the CD11c-bright, highly autofluorescent population and do not express MHCII. DCs are identified as CD11c-bright, low autofluorescent cells, which strongly express MHCII. DCs enumerated by these criteria correspond with myeloid DCs. The following antibodies were used to stain mouse T-cell subpopulations: fluorescein isothiocyanate–conjugated anti-CD4 (L3T4), phycoerythrin-conjugated anti-CD8 (Ly-2), and APC-conjugated anti-CD3 (145-2C11) monoclonal antibodies. All monoclonal antibodies were obtained from BD Pharmingen (San Diego, CA).
Cells were incubated with 7-amino-actinomycin (7-AAD or viaprobe; BD Pharmingen) for dead cell exclusion. All labeling reactions were performed on ice in FACS-EDTA buffer.
Flow cytometry data acquisition was performed on a dual-laser FACS VantageTM flow cytometer running CELLQuest software (Becton Dickinson, Mountain View, CA). FlowJo software (www.Treestar.com) was used for data analysis.
To obtain lung tissue, right heart catheterization and perfusion with saline-EDTA was performed to remove the pulmonary intravascular pool of cells. The right lung was clamped, removed, and collected in ice-cold tissue culture medium for preparation of single-cell suspensions. Lung macrophages were isolated as described previously (27). To obtain sufficient amounts of macrophages for RNA preparation, the single-cell suspensions of eight mice were pooled. Pulmonary single-cell suspensions were first incubated with Fc-receptor-block, followed by anti-CD11c microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). CD11c+ lung cells were enriched after one passage through a VarioMACS magnetic cell separator according to the manufacturer's instructions. The CD11c− fraction was kept on ice until RNA extraction. The CD11c+ fraction was labeled with APC-conjugated anti-CD11c. On the FACS Vantage SE, the CD11c+/highly autofluorescent cells, representing the macrophage population, were separated by fluorescence-activated cell sorting with a 70-nm nozzle.
The left lung was fixated by infusion of 4% paraformaldehyde through the tracheal cannula (22–24). After excision, the lung was immersed in fresh fixative for 2 hours. The lung lobe was embedded in paraffin, cut into 3-μm transverse sections, and stained for histologic analysis.
Histolocalization of HA was determined on paraffin sections using biotin-labeled HA-binding protein (HABP-b) (Seikagaku, Tokyo, Japan). Sections were subjected to deparaffinization followed by rehydration. Sections were stained with HABP-b (2 μg/ml) at room temperature for 1 hour. After washing, the DAKO Cytomation Streptavidine ABComplex/HRP system was used according to the manufacturer's instructions (DAKO, Glostrup, Denmark). Enzymatic reactivity was visualized with the Vector NovaRED peroxidase substrate kit (Vector, Burlingame, CA). Sections were lightly counterstained with hematoxylin and mounted in Faramount (DAKO). No significant staining was detected in sections pretreated with 50 U/ml Streptomyces hyaluronidase (Calbiochem, San Diego, CA) at 37°C for 2 hours, indicating that this HABP staining reaction was specific for HA.
The HABP-b staining in the alveolar walls was semiquantitatively scored by three independent observers (J.C., R.V.S., and M.D.) who were unaware of the treatment of the animals. The intensity of the HA staining was expressed in arbitrary units and scored on a five-point scale: 0 = no or hardly any staining, 1 = weak staining at some spots on the alveolar walls, 2 = moderate staining at several spots, 3 = intense staining of most parts of the alveolar walls, and 4 = very intense staining throughout the alveolar walls.
The HABP-b staining in the airway walls was scored quantitatively, and three lung sections per animal were examined. The following morphometric parameters (28) were marked manually on the digital representation of the airway: the length of the basement membrane (Pbm), the area defined by the basement membrane (Abm), and the area defined by the total adventitial perimeter (Ao). The total bronchial wall area (WAt) was calculated (WAt = Ao – Abm) and normalized to the squared length of the basement membrane. For the quantification of HA deposition, the area in the airway wall covered by the stain was determined by the software (KS400; Zeiss, Oberkochen, Germany), and its value was calculated as described previously (29). The area of HA was normalized to Pbm. All airways with a Pbm smaller than 2,000 μm and cut in reasonable cross-sections (defined by a ratio of minimal to maximal internal diameter >0.5) were included.
Collagen in the airway wall was stained using Sirius Red, and the amount of fibronectin was determined with a goat antirat fibronectin antibody (Calbiochem, BadsSoden, Germany) using the streptavidin-biotin peroxidase method as previously described (24, 29). Bronchial collagen and fibronectin staining was scored quantitatively as described previously for HABP-b staining.
Emphysema is a structural disorder characterized by destruction of the alveolar walls and enlargement of the alveolar spaces. We determined enlargement of alveolar spaces by quantifying the mean linear intercept (Lm) after 24 weeks of smoke exposure as described previously (22, 30) using image analysis software (Image J 1.33). Only sections without cutting artifacts, compression, or hilar structures (airway or blood vessel with a diameter larger than 50 μm) were used in the analysis. The Lm was measured by placing a 100 × 100 μm grid over each field. The total length of each line of the grid divided by the number of alveolar intercepts gives the average distance between alveolated surfaces, or the Lm.
Lung tissue specimens were homogenized by a Polytron homogenizer (Kinematica, Luzern, Switserland) (5 × 10-s bursts with 1-min intervals in ice) in 10 ml of 25 mM Tris-HCl (pH 7.6) per gram of tissue. Homogenized tissues were delapidated with 4 volumes of chloroform/methanol (1:2 volumes). Organic solvents were removed by centrifugation (3,(200 × g, 20 min, 4°C), and the pellet was washed with 10 ml of ethanol, centrifuged as described previously, and dried at 40°C for 4 hours. The dry pellet was weighed, resuspended in 1 ml of 0.1 M Tris-HCl buffer (pH 8.0) containing 1 mM CaCl2, and subjected to protein digestion with 0.1 KU of Pronase (Streptomyces griseus; Calbiochem). The pronase solution was preincubated for 30 minutes at 60°C to eliminate glycosidase activity. Digestion was performed for 72 hours at 60°C by adding equal amounts of pronase at 24-hour intervals. The sample concentration was then adjusted to 150 mM NaCl and 10 mM MgCl2, and DNA digestion was accomplished by adding 400 KU DNase I (EC 3.1.21.1; Calbiochem) and incubating for 16 hours at 37°C. At the end of the incubation period, the CaCl2 concentration of the solution was adjusted to 1 mM, and the reaction was stopped by adding 0.1 KU of pronase and incubating the mixture at 60°C for 24 hours. The pH was adjusted to 10.0 to 11.0 by the addition of 10 mM NaOH, and samples were subjected to β elimination in the presence of 1 M NaBH4 for 16 hours at 45°C. Samples were then neutralized with 50% (vol/vol) acetic acid. Total glycosaminoglycans were precipitated with the addition of four volumes of ethanol in the presence of 0.1 volume of 3 M CH3COONa and overnight maintenance at −4°C, recovered with centrifugation (20 min; 2,000 × g), redissolved in double distilled H2O, and stored at 4°C. Colorimetric determination of uronic acids was performed according to Bitter and Muir (31).
The relative amount of HA was measured in aliquots containing 0.075 μg of uronic acids by ELISA (Corgenix, Lynch Wood, Peterborough, UK) as previously described (32). Briefly, ELISA plates coated with HA-binding protein were incubated with samples or standards (1 h at room temperature) in duplicates, washed five times with washing buffer, incubated with a solution containing horseradish peroxidase–conjugated HA-binding protein (30 min at room temperature), washed again five times, and incubated with 100 μl of the substrate solution. After 30 minutes, the reaction was stopped by adding an equal amount of sulfuric acid (0.36 N), and the optical density was measured at 450 nm (630-nm reference).
Total glycosaminoglycans (6 μg of uronic acids), isolated and purified from lung tissue specimens, were analyzed on 0.9% agarose gels, as previously described (33). Identification of HA was performed by enzymatic treatment of the samples with hyaluronidase (Streptomyces hyalurolyticus; Seikagaku, Tokyo, Japan) for 14 hours at 40°C before gel electrophoresis. The molecular mass of HA was determined using commercially available HA of 1,000, 250, and 50 kD (AMS Biotechnology, Oxfordshire, UK). Gels were stained with a solution of 0.005% (wt/vol) stains-all dissolved in 50% (vol/vol) ethanol overnight under light-protective cover at room temperature. For destaining, gels were transferred in H2O and incubated for 3 hours at room temperature.
Total lung RNA and RNA from isolated lung macrophages was extracted with the RNeasy Mini Kit (Qiagen, Hilden, Germany). cDNA was obtained by reverse transcription of RNA using a 3:1 ratio of hexa/oligodT primers (Abgene, Epsom, UK). The primer pairs used in this study were designed with Primer Express 2.0 Software (Applied Biosystems, Foster City, CA) and are listed in Table 1. Real-time PCR reactions were performed in duplicate using diluted cDNA template (dilution 1:25), 0.6 pmol of each primer (Table 1), and IQ SYBR Green Supermix I dye (Bio-Rad, Hercules, CA) in a total volume of 20 μl. A standard curve derived from the serial dilutions of a mixture of all samples was included on each plate. Quantitation and real-time detection of the PCR products was followed on an iCycler iQ Real-Time PCR system (Bio-Rad) with the following cycling conditions: 3 minutes at 95°C for Platinum Taq activation and 40 cycles for the melting (15 s at 95°C) and annealing/extension (45 s, 60°C) steps. These conditions generate specific PCR products of the desired length, which were verified by gel electrophoresis on an ethidium bromide–stained 2% agarose gel. Data were processed using the standard curve–based method. Expression of target genes was corrected by a normalization factor that was calculated based on expression of three reference genes (Hprt1, Ppia, and Rpl13a) using the geNorm applet according to the guidelines and theoretical framework previously described (http://medgen.ugent.be/∼jvdesomp/genorm/) (34).
Gene | Accession No. | Sequences | Product |
|---|---|---|---|
| Has1 | NM_008215 | Forward: 5′-ACCTCACCAACCGAATGCTT-3′ | 89 bp |
| Reverse: 5″-GAAGGAAGGAGGAGGGCG-3′ | |||
| Has2 | NM_008216 | Forward: 5′-TGAGTACAAAGAGGTTCGTTCAAGTT-3′ | 86 bp |
| Reverse: 5′-ATTGTCAGGGTGTGTTTGTTTCC-3′ | |||
| Has3 | NM_008217 | Forward: 5′-CTACTTTGTAGCTGCCCAGAATACTG-3′ | 116 bp |
| Reverse: 5′-GAGTACAAAAAACAGCACCGGAAT-3′ | |||
| Hyal1 | NM_008317 | Forward: 5′-CTTCTGCCCCTGGAGGAACT-3′ | 141 bp |
| Reverse: 5′-GTGTGGAATCCATGTATGCTTTAATG-3′ | |||
| Hyal2 | NM_010489 | Forward: 5′-CGAGGACTCACGGGACTGA-3′ | 150 bp |
| Reverse: 5′-GCTGAGTTAGGTAATTCTTGAGGTATTG-3′ | |||
| Rpl13a | NM_009438 | Forward: 5′-CACTCTGGAGGAGAAACGGAAGG-3′ | 182 bp |
| Reverse: 5′-GCAGGCATGAGGCAAACAGTC-3′ | |||
| Ppia | NM_008907 | Forward: 5′-TTCCTCCTTTCACAGAATTATTCCA-3′ | 75 bp |
| Reverse: 5′-CCGCCAGTGCCATTATGG-3′ | |||
| Hprt1 | NM_013556 | Forward: 5′-AGCTACTGTAATGATCAGTCAACG-3′ | 198 bp |
| Reverse: 5′-AGAGGTCCTTTTCACCAGCA-3′ |
Reported values are expressed as mean ± SEM. Statistical analysis was performed with Sigma Stat software (SPSS 11.0 Inc., Chicago, IL) using nonparametric tests (Kruskall-Wallis and Mann-Whitney U). P values under 0.05 were considered as significant.
Upon subacute (4-wk) and chronic (24-wk) exposure to CS, the total cell number in the BAL fluid was significantly increased, compared with air-exposed control animals (Figure 1A). Exposure to CS caused a significant accumulation of macrophages, DCs, neutrophils, CD4+ T lymphocytes, and CD8+ T lymphocytes in the BAL fluid (Figures 1B–1F).
Figure 1. Effect of cigarette smoke-exposure on cell differentiation in bronchoalveolar lavage (BAL) fluid. Total BAL cells and cell differentiation in BAL fluid upon subacute (4-wk) and chronic (24-wk) exposure to air or cigarette smoke. (A) Total BAL cells. (B) Macrophages. (C) Dendritic cells. (D) Neutrophils. (E) CD4+ T lymphocytes. (F) CD8+ T lymphocytes. Results are expressed as means ± SEM; n = 8 animals per group (*P < 0.05).
[More] [Minimize]Histochemical localization using HA binding protein revealed little HA staining in alveolar walls of air-exposed mice. Semiquantitative scoring of the HA staining in alveolar walls showed only slightly enhanced HA deposition upon subacute (4-wk) CS exposure, whereas chronic (24-wk) CS exposure caused a significant increase in the deposition of HA (Figure 2).
Figure 2. Effect of cigarette smoke-exposure on hyaluronan (HA) deposition in alveolar walls. (A) Semiquantitative scoring of HA deposition in alveolar walls upon subacute (4-wk) and chronic (24-wk) exposure to air or cigarette smoke (CS). Results are expressed as means ± SEM; n = 8 animals per group (*P < 0.05). (B–E) Photomicrographs of HA deposition in alveolar walls upon sub-acute (4-wk) and chronic (24-wk) air or CS exposure. (B) Air exposure, 4 wk. (C) CS exposure 4 wk. (D) Air exposure, 24 wk. (E) CS exposure, 24 wk.
[More] [Minimize]Histochemical localization of HA revealed constitutive staining in airway walls of air-exposed mice. Quantitative scoring using an image analyzer showed that subacute (4-wk) and chronic (24-wk) exposure to CS caused a significant increase in peribronchial deposition of HA compared with air-exposed animals (Figure 3).
Figure 3. Effect of CS exposure on peribronchial HA deposition. (A) Quantitative scoring of peribronchial HA deposition upon sub-acute (4-wk) and chronic (24-wk) exposure to air or CS. Results are expressed as means ± SEM; n = 8 animals per group (*P < 0.05). Pbm = length of basement membrane. (B–E) Photomicrographs of peribronchial HA deposition upon subacute (4-wk) and chronic (24-wk) air or CS exposure. (B) Air exposure, 4 wk. (C) CS exposure, 4 wk. (D) Air exposure, 24 wk. (E) CS exposure, 24 wk.
[More] [Minimize]Peribronchial deposition of protein components of the ECM was studied on Sirius Red– and antifibronectin-stained histologic sections to reveal collagen in fibronectin, respectively. Subacute (4-wk) exposure to CS did not induce an increase in the peribronchial deposition of collagen and fibronectin (Figure 4). Upon chronic (24-wk) exposure to CS, there was a significant increase in deposition of collagen and fibronectin in airway walls as compared with air-exposed control mice (Figure 4).
Figure 4. Effect of CS exposure on peribronchial collagen and fibronectin deposition. (A) Quantitative scoring of peribronchial collagen and (F) fibronectin deposition upon subacute (4-wk) and chronic (24-wk) exposure to air or CS. Results are expressed as means ± SEM; n = 8 animals per group (*P < 0.05). Pbm = length of basement membrane. (B–J) Photomicrographs of peribronchial collagen and fibronectin deposition upon subacute (4-wk) and chronic (24-wk) air or CS exposure. (B, G) Air exposure, 4 wk. (C, H) CS exposure, 4 wk. (D, I) Air exposure, 24 wk. (E, J) CS exposure 24 wk.
[More] [Minimize]Pulmonary emphysema is characterized by the destruction of alveolar walls due to damage to the lung parenchyma, leading to enlargement of alveolar spaces. Therefore, we quantified emphysematous lesions by measuring the mean linear intercept (Lm). Chronic (24-wk) exposure to CS clearly induced pulmonary emphysema in wild-type animals, as evidenced by a significant increase in the Lm (air 38.0 ± 0.9 μm versus CS 41.9 ± 0.5 μm; P < 0.05).
Total glycosaminoglycans were isolated and purified from lung tissue specimens. Measurement of the content of uronic acids in lung samples revealed that they contain 7.0 ± 0.8 μg of uronic acids per mg of dry defatted lung tissue. There were no significant differences in the content of total glycosaminoglycans in dry defatted lung tissue between the different groups tested (data not shown).
The content of HA in total glycosaminoglycans isolated and purified form lung tissue specimens was measured in 0.075 μg of uronic acids using ELISA. There were no significant differences in the relative content of HA in total glycosaminoglycans in the lung of subacute (4-wk) CS-exposed mice. However, we observed a significant increase of the relative content of HA after chronic (24-wk) CS exposure (Figure 5A).
Figure 5. Effect of CS exposure on the relative amount and molecular mass of HA in the lung. Total glycosaminoglycans were isolated and purified from the lungs upon subacute (4-wk) and chronic (24-wk) exposure to air or CS. (A) The relative amount of HA in aliquots of total glycosaminoglycans containing 0.075 μg of uronic acids was determined by ELISA. Results are expressed as means ± SEM; n = 8 animals per group (*P < 0.05). (B) Representative results obtained after electrophoresis of total glycosaminoglycans corresponding to 6 μg of uronic acids on 0.9% agarose gels. HA (1 μg) of known molecular mass was used as a marker. Gels were stained with a solution of 0.005% (wt/vol) stains-all dissolved in 50% (vol/vol) ethanol and destained in H2O.
[More] [Minimize]We analyzed the molecular mass of HA that was isolated and purified from lung tissue specimens by agarose gel electrophoresis. HA isolated from the lungs of air-exposed mice migrated as a broad band, with an average molecular mass of 500 kD. HA isolated from the lungs of CS-exposed mice exhibited a lower molecular mass of an average of 70 kD after subacute (4-wk) and chronic (24-wk) CS exposure (Figure 5B).
Expression of HA synthase genes and hyaluronidases was studied on total lung tissue by RT-PCR. Subacute (4-wk) and chronic (24-wk) CS exposure significantly increased the mRNA expression of Has3 compared with air-exposed mice (Figure 6C). There was no effect of CS exposure on the expression of Has2 (Figure 6B), whereas chronic exposure to CS significantly decreased the mRNA expression of Has1 compared with air-exposed control mice (Figure 6A).
Figure 6. Effect of CS exposure on the expression of HA-modulating genes in total lung tissue mRNA levels of (A) Has1, (B) Has2, (C) Has3, (D) Hyal1, and (E) Hyal2 upon subacute (4-wk) and chronic (24-wk) exposure to air or CS as measured by RT-PCR. mRNA levels were corrected using a Genorm calculated normalization factor based on mRNA expression of three reference genes (Rpl13a, Ppia, and Hprt1). Results are expressed as means ± SEM; n = 5 animals per group (*P < 0.05). Hprt1 = hypoxanthine phosphoribosyltransferase 1; Ppia = cyclophilin A/peptidylprolyl isomerase A; Rpl13a = ribosomal protein L13a.
[More] [Minimize]The mRNA expression of Hyal1 was not affected by CS exposure (Figure 6D). However, there was a tendency toward decreased mRNA expression of Hyal2 upon chronic CS exposure compared with air-exposed mice (Figure 6E).
Expression of HA synthase genes and hyaluronidases was studied by RT-PCR in macrophages isolated from lungs of air- and CS-exposed mice. Upon subacute CS exposure, there was a strong decrease in Has1 mRNA in lung macrophages (Figure 7A), whereas Has2 and Has3 were only slightly decreased (Figures 7B–7C). In contrast to the HA-synthase genes, the hyaluronidase Hyal2 was strongly increased in macrophages from CS-exposed mice (Figure 7E) compared with macrophages from air-exposed mice. Hyal1 mRNA expression was not affected by CS exposure (Figure 7D).
Figure 7. Effect of CS exposure on the expression of HA-modulating genes in isolated lung macrophages mRNA levels of (A) Has1, (B) Has2, (C) Has3, (D) Hyal1, and (E) Hyal2 in macrophages isolated from lung tissue upon subacute (4-wk) exposure to air or CS as measured by RT-PCR. mRNA levels were corrected using a Genorm calculated normalization factor based on mRNA expression of three reference genes (Rpl13a, Ppia, Hprt1). Results are expressed as means ± SEM; n = 2 separate experiments. Hprt1: hypoxanthine phosphoribosyltransferase 1; Ppia = cyclophilin A/peptidylprolyl isomerase A; Rpl13a = ribosomal protein L13a.
[More] [Minimize]Exposure to CS is the main risk factor for the development of COPD, a disease that is characterized by chronic inflammation, destruction of lung parenchyma, and airway wall remodeling. In this study we show in a murine model of COPD that CS-induced pulmonary inflammation is accompanied by enhanced deposition of HA in alveolar and bronchial walls. We observed an increase in proinflammatory, LMW, HA fragments in lungs of CS-exposed mice. The increased HA deposition is most likely caused by CS-induced modulation of HA synthase and hyaluronidase genes.
The increased deposition of HA in bronchial walls of CS-exposed mice may contribute to the airway wall remodeling that is a hallmark of COPD pathology and leads to airway obstruction. Similarly, we have recently described that increased peribronchial deposition of collagen and fibronectin upon chronic CS exposure results in thickening of the airway walls in mice (23, 24). The levels of the glycosaminoglycan HA were already increased upon subacute CS exposure, whereas the deposition of collagen and fibronection (protein components of the ECM) did not increase before chronic CS exposure. Although chronic CS exposure also led to destruction of alveolar walls or emphysema, there was increased deposition of HA in the remainder of the alveolar walls.
The enhanced HA content in bronchial and alveolar walls is most likely the result of a skewed synthesis and breakdown balance, as can be concluded from the increased expression of the HA synthase Has3 and the decreased expression of Has1 in total lung tissue. Because Has3 is known to produce LMW HA fragments, whereas Has1 creates HMW HA (20), their regulation upon CS exposure may result in an accumulation of proinflammatory LMW HA fragments. Indeed, we observed a shift in the molecular weight of HA from HMW HA in lungs of air-exposed mice toward LMW HA fragments in lungs of CS-exposed mice. Degradation of HA and reduced chain length of HA in lungs of patients with emphysema has been reported in older literature (35, 36). LMW HA fragments can act as endogeneous danger signals (so-called “damage associated molecular patterns” or DAMPs) by binding to CD44, Toll-like receptor (TLR)2, or TLR4. That way they contribute not only to the initiation of airway inflammation but also to the persistence of chronic airway inflammation, even after smoking cessation. It has been shown that LMW HA fragments can induce macrophages to release the CCR5 chemokines MIP-1α/CCL3, MIP-1β/CCL4, and RANTES/CCL5 and the CCR2 chemokine MCP-1/CCL2 (17). These chemokines have been found in increased levels in lungs of CS-exposed mice and in patients with COPD (37). Moreover, the chemokine receptor CCR5 has recently been implicated in the pathogenesis of COPD (24, 38). Additionally, LMW HA fragments can stimulate macrophages to release matrix metalloproteinase–12 (18), a proteolytic enzyme that has been described as one of the key players in the development of CS-induced emphysema in mice (39). In contrast, opposite biological functions have been described for HMW HA. Indeed, HMW HA has antiinflammatory effects by, for example, suppressing chemokine expression (40), and it is also involved in tissue repair (41). Cantor and colleagues reported protective effects of aerosolized HA on chronic CS-induced lung injury and emphysema (42).
To elucidate a cellular source of the HA-modulating enzymes, we studied their expression in in vivo isolated lung macrophages. Consisting with the expression in total lung tissue, Has1 was down-regulated in macrophages from CS-exposed mice. In contrast to total lung tissue, the expression of Hyal2 was strongly up-regulated in macrophages from CS-exposed mice. This may point to a role of macrophages in the clearance of excessive HA and in the breakdown of HMW HA to LMW HA fragments. The expression of HA-modulating genes needs to be examined in other cellular sources, such as bronchial epithelial cells or pneumocytes, to completely understand the levels that are found in total lung tissue.
It has been reported that several cytokines (e.g., TNF-α, IFN-γ, and IL-1β) and growth factors (e.g., TGF-β), but also oxidative stress, can stimulate the production of HA by altering the expression of HA-modulating enzymes (43–46). We examined the role of TNF-α in CS-induced HA deposition by subjecting TNF-α R1 and TNF-α R2 knockout (KO) mice to chronic CS exposure. We could not detect any differences between wild-type and TNF-α R1 or TNF-α R2 KO mice in HA deposition or in expression of HA-modulating genes (data not shown). However, the involvement of TNF-α cannot be excluded because we did not use TNF-α R1 and TNF-α R2 double KOs or a TNF-α blocking antibody.
HA exerts many of its biological functions through the CD44 receptor that is expressed on many cell types, including leukocytes, epithelial cells, endothelial cells, and fibroblasts (47). The binding of HA to CD44 is regulated by T-cell– and monocyte-derived cytokines, primarily TNF-α (48). HA can also bind to other receptors, including TLRs. Oligosaccharides of HA have the capacity to activate DCs through TLR4 (16, 49). Therefore, it is tempting to speculate that the increased LMW HA content in lungs of CS-exposed mice can contribute to the previously described CS-induced maturation of DCs (22, 50). Moreover, this DC maturation was significantly impaired in CS-exposed, TLR4-deficient mice. HA can also influence the function of regulatory T cells. Indeed, HMW HA conveys antiinflammatory signals by promoting the suppressive effects of regulatory T cells (51).
Although in vivo CS-exposed mice can offer valuable information on several aspects of the pathogenesis COPD, such as the time course of pulmonary inflammation, there are limitations that need to be taken into account. First, there are certain anatomic and physiologic differences between the respiratory tract of mice and humans. For example, mice are obligate nose breathers that filter tobacco smoke inefficiently, and they have less branching of the bronchial tree. Second, the profile of inflammatory mediators is slightly different in the mouse. Third, there is no mouse model that mimics all the hallmarks of COPD pathology, including exacerbations and extrathoracic manifestations. As a result of these limitations, further studies on human material are needed and are in progress.
In summary, we showed for the first time that HA deposition is increased in alveolar and bronchial walls of CS-exposed mice. We provided evidence that CS exposure leads to increased accumulation of LMW, proinflammatory HA fragments. These findings correlated with CS-induced changes in the expression of HA-modulating genes. Moreover, we found that increased deposition of HA already occurs upon subacute CS exposure, which is much earlier than the deposition of fibronectin or collagen. This increase in HA may contribute not only to airway wall remodeling, but also to the ongoing pulmonary inflammation in COPD. However, the molecular and cellular mechanisms that lead to this exaggerated deposition of HA need to be elucidated.
The authors thank Greet Barbier, Eliane Castrique, Indra De Borle, Philippe De Gryze, Katleen De Saedeleer, Anouck Goethals, Johanna Jörgensen, Marie-Rose Mouton, Ann Neessen, Christelle Snauwaert, and Evelyn Spruyt for their technical assistance.
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