Abstract
Hyaluronan, a linear polysaccharide, is accumulated in lung interstitium during different pathological conditions, causing interstitial edema and thereby impaired lung function. We investigated the mechanism of local hyaluronan turnover during the early phase of bleomycin-induced fibrotic lung injury in rats. The binding of [3H]hyaluronan to alveolar macrophages (AM) established from bleomycin-treated rats 1 and 5 days after induction of injury was decreased 8- and 15-fold, respectively, compared with that of AM from saline-treated control counterparts, but at day 14 returned almost to the normal level. Data was confirmed by quantitative cytochemistry, using fluorescein-labeled hyaluronan. Analysis of the expression of CD44, a receptor for hyaluronan, by Western blotting revealed a 30% increase of CD44 molecules expressed on AM from bleomycin-treated rats at day 5 compared with control rats. In particular a lower molecular mass form of CD44 appeared. No expression of the receptor for hyaluronan-mediated motility (RHAMM) could be detected. The internalization and degradation of [3H]hyaluronan by AM, obtained from bleomycin-treated rats at days 1, 5, and 14, were decreased about 65%, 35%, and 30%, respectively, compared with AM from the control rats. The AM lysosomal hyaluronidase activity did not differ significantly between bleomycin-treated and control rats. Our results indicate that a decreased hyaluronan binding capacity of AM may account for the impairment of internalization and thereby degradation of excessive hyaluronan during the early phase of fibrotic lung injury.
Hyaluronan, a high molecular weight polysaccharide, is present in different amounts in all body tissues, and it has been suggested to carry important functions during embryogenesis, cell growth and migration, angiogenesis, inflammation and wound healing (for review, see reference 1). Interactions between hyaluronan and its binding receptors, CD44 and receptor for hyaluronan-mediated motility (RHAMM), are important for the biological effects of polysaccharide (2). Most of the synthesized hyaluronan is degraded at the site of origin and during filtration through lymph nodes (3). Alveolar macrophages (AM) are the major cell type known to internalize hyaluronan for degradation in normal lungs (4, 5). AM possess specific hyaluronan receptors, CD44 molecules, which are the most extensively studied representatives of hyaladherins (for review, see reference 6). CD44 is a widely distributed family of cell surface glycoproteins with a variety of functions (7), but not all cells that express CD44 are able to bind hyaluronan. Hyaluronan binding to CD44 can be dependent on different functional states of receptor bearing cells (8), or cell type-specific glycosylation of the receptor (9, 10). After ligation and internalization the hyaluronan is subject to specific enzymatic digestion (11). Part of the hyaluronan is drained into the lymph (12) and catabolized or fragmented in lymph nodes (13). Only the minor part with average Mr about 2 × 105 is reaching circulation and is rapidly degraded by liver endothelial cells (14). In lung disorders with hyaluronan overproduction the above-described mechanisms fail to maintain equilibrium, which results in hyaluronan accumulation in the lung interstitium.
The early phase of fibrotic lung injury is characterized by transient deposition of hyaluronan in the lung interstitium caused by proliferation (15) and activation (16-18) of fibroblasts, which is paralleled by water accumulation and influx of inflammatory cells (19, 20). The resulting severe architectural changes of the alveolar walls cause the loss of functional alveolar capillary units which may lead to respiratory insufficiency. Although the characteristic changes have been described and hyaluronan in bronchoalveolar lavage fluid (BALF) has been suggested to be a sensitive marker of lung injury, the underlying molecular mechanisms of hyaluronan accumulation are not completly understood.
The primary aim of the present study was to further determine the mechanisms involved in the accumulation of hyaluronan in the early phase of fibrotic lung injury, namely the local turnover of hyaluronan in bleomycin-treated rats.
RPMI 1640 medium and phosphate-buffered saline without Ca2+ and Mg2+ (PBS) were purchased from Swedish Veterinary Institute (Uppsala, Sweden). Tenfold concentrated PBS was purchased from Gibco Life Technologies AB (Täby, Sweden). Percoll was bought from Pharmacia Biotech AB (Uppsala, Sweden). Radioactive hyaluronan, labeled with 3H in the acetyl groups ([3H]hyaluronan, Mr 0.97 × 106, 0.2 μCi/μg) was a generous gift from Dr. J. R. E. Fraser (Melbourne, Australia). Hyaluronan of Mr 1.3 × 106 was kindly provided by O. Wik (Pharmacia, Uppsala, Sweden). DNase I was purchased from Boehringer Mannheim (Mannheim, Germany). Agarose SeaKem was purchased from FMC BioProducts (Rockland, ME). Testicular hyaluronidase (360 units per mg protein) was obtained from Sigma (St. Louis, MO). Monoclonal antibodies ED1 to rat lung macrophages were kindly provided by Dr. G. Kraal (Amsterdam, The Netherlands). Monoclonal anti-rat CD44 antibodies 5G8 (anti exon 15) and 1.1ASML (anti-CD44v6) were a generous gift from Drs. J. P. Sleeman and P. Herrlich (Karlsruhe, Germany). Polyclonal antibodies R 3.4 against peptides encoded in the RHAMM cDNA were kindly provided by Drs. R. Savani and E. Turley (Winnipeg, Manitoba, Canada).
The study was approved by the Ethical Committee for Laboratory Animals in Uppsala. Adult male Sprague-Dawley inbred rats (ALAB, Sollentuna, Sweden) weighing about 200 g were used. Animals were kept in separate cages, and provided with standard chow and tap water ad libitum. For our study we used a model of fibrotic lung injury in rats, initially described by Thrall and coworkers (21). Briefly, 1.5 IE bleomycin sulphate (Lundbeck, Copenhagen, Denmark) in 0.3 ml sterile saline was administrated to the animals through single intratracheal injection under chloralhydrate-pentobarbital anesthesia. Control animals received 0.3 ml of sterile saline. All rats received 10 mg sodium cefuroxim (Glaxo, Greenford, Middlesex, UK) intraperitoneally immediately after tracheal cannulation in order to minimize the risk of bacterial infections.
The procedure for AM isolation was modified from Lavnikova and colleagues (22). The animals were exsanguinated under chloralhydrate-pentobarbital anesthesia 1, 5, or 14 days after intratracheal injection. Vascular perfusion with 25 ml of PBS containing 0.6 mM EDTA (PBS-EDTA) at 37°C was performed via the right heart.
Using an in situ technique, lungs were lavaged by intratracheal infusion of 6 aliquots each of 8 ml of PBS-EDTA at 37°C at a hydrostatic pressure of 15 cm. After 5 min, the fluid was recovered by gravity and massage of the thorax into polypropylene tubes kept on ice. The BALF was centrifuged at 820 × g for 10 min at 4°C and pellet was dissolved in 30 ml of PBS containing 0.3% bovine serum albumin (BSA), 100 U/ml penicillin, 100 μg/ml streptomycin, and 1 μg/ml fungizone (PBS-BSA).
Following vascular perfusion and repeated bronchoalveolar lavage with PBS-EDTA, the trachea, major bronchi, and large blood vessels were removed and the lung tissue was minced to about 5 mm3 fragments followed by continuous end-over-end agitation in 40 ml PBS containing 50 μg/ml DNase I for 30 min at 4°C. The dispersed lung cells were filtrated through a 53-μm sterile nylon mesh.
Both BALF- and tissue-AM were processed in the same manner at 4°C. The tubes were shaken at 150 rpm for 30 min on a rocking platform (New Brunswick Scientific Co., Edison, NJ) and then centrifuged at 820 × g for 10 min followed by washing under same conditions with 30 ml of PBS-BSA. In order to separate AM from lymphocytes, granulocytes, erythrocytes and debris, the pellets were resuspended in 3 ml of PBS-BSA and mixed with 7 ml of 100% Percoll (9 parts of Percoll diluted with 1 part of 10-fold concentrated PBS). The cell suspensions were underlayered a two-step Percoll gradient (15 ml of 65% Percoll, 20 ml of 35% Percoll, and 5 ml of PBS-BSA) in 50-ml Falcon tubes, and centrifuged at 1,850 × g for 30 min without brakes (Figure 1). Three different cell bands were obtained. Debris was found on the top of the Percoll gradient (fraction 1), and erythrocytes, lymphocytes, and granulocytes were pelleted (fraction 3). Both these fractions were discharged. AM were obtained from the interface between 65% and 35% Percoll layers (fraction 2) and resuspended in 50 ml of PBS-BSA followed by centrifugation at 820 × g, 10 min, in order to remove the Percoll solution. The pellets were resupended in PBS-BSA and total cell numbers were determined in Bürker chambers followed by cytospin analysis for quantification of AM. The average numbers of BALF- and tissue-AM from control rats at 95–99% purity were 6.9 × 106 and 3.1 × 106 cells, respectively, and 6.2 × 106 and 6.5 × 106 cells from bleomycin-treated rats, respectively. After centrifugation at 820 × g, 10 min, the pellets were resuspended in 37°C RPMI 1640 medium containing 100 U/ml penicillin, 100 μg/ml streptomycin, 4 mM L-glutamine (1 × 106 AM per ml). Then known amounts of AM were seeded into the cell culture plates according to experimental setup and incubated at 37°C, 5% CO2 in a humidified atmosphere for 10 min. Non-adherent cells (about 1–3% of total cell numbers, mainly granulocytes) were removed by replacement of media. The plates, which after such treatment contained only AM (23), were examined with a light microscope using phase contrast optics (Leitz, Wetzlar, Germany) and quantified by staining with ED1 antibodies recognizing all types of macrophages.
Fig. 1. Purification of rat AM by two-step Percoll gradient. Schematic illustration of the purification of BALF- and tissue-AM from rats, as described in Materials and Methods.
[More] [Minimize]Hyaluronan from Pharmacia Biotech AB, Uppsala, Sweden (Mr 1.3 × 106) was labeled with fluoresceinamine isomer I (Sigma) after cyanogen bromide activation of the polysaccharide by the method of Glabe (24). All steps were carried out at room temperature. Briefly, 20 mg of hyaluronan were activated at pH 11 by adding 10 mg of cyanogen bromide in 10 mM sodium perborate for 5 min. The activated polysaccharide was separated on a Sephadex G-25 column (PD-10; Pharmacia, Uppsala, Sweden) equilibrated with 0.2 M borate buffer at pH 8 and then incubated at continuous agitation overnight with 2 mg fluoresceinamine. Unbound fluoresceinamine was separated from hyaluronan-bound fluoresceinamine on a PD-10 column equilibrated with PBS at pH 7.5. Hyaluronan concentration was determined with a commercial kit Test-50 (Pharmacia, Uppsala, Sweden) and fluorescence (exitation at 492 nm and emission at 514 nm) on a Hitachi F4000 fluorescence spectrophotometer.
The binding assay of [3H]hyaluronan was performed essentially as described by Asplund and Heldin (25). Subconfluent AM cultures (2 × 105 cells per well, 24-well plate) were cultured for 24 h at 37°C in 5% CO2 in serum-free RPMI 1640 medium. Under such conditions testicular hyaluronidase treatment did not increase the hyaluronan binding to AM, and was skipped (data not shown). Then, fresh medium (4°C) containing increasing concentrations of [3H]hyaluronan (0 to 2 μg/ml) was added in the absence or presence of non-labeled hyaluronan (100 μg/ml; Mr 1.3 × 106) in a final volume of 300 μl. Following incubation for 24 h at 4°C the unbound [3H]hyaluronan was removed by three washes with ice-cold RPMI 1640 medium. The cell layers were then dissolved in 300 μl of 0.3 M NaOH containing 1% sodium dodecyl sulfate (SDS) for 30 min at room temperature on a rocking platform. Fifty μl of 2 M HCl and 2 ml of Ecoscint A scintillation fluid (Hintze, Lidingö, Sweden) were then added to lysates and the samples were subjected to scintillation counting. The radioactivity was determined in a Pharmacia LKB Wallac scintillation counter. Specific binding was calculated by subtraction of the background label which was measured in the presence of excess of non-labeled hyaluronan.
The binding assay of fluorescein-labeled hyaluronan was performed with conventional technique. Briefly, 2 × 105 cells were cultured on coverslips in RPMI 1640 medium for 24 h at 37°C in 5% CO2. After gentle washing, the cells were incubated with fluorescein-labeled hyaluronan (100 μg/ml, Mr 1.3 × 106) in RPMI 1640 in absence or presence of unlabeled hyaluronan (500 μg/ml, Mr 1.3 × 106) for 16 h at 4°C. Following three washes with ice-cold PBS, the cells were fixed in 100% ice-cold acetone for 10 min, air-dried, and then rehydrated 15 min in PBS. Processed cells were mounted in Fluorimount, and examined with a light microscope equipped with fluorescence optics (Nikon, Japan).
Subconfluent AM cultures (3 × 106 cells per 35-mm dish) were cultured in serum-free RPMI 1640 medium for 24 h at 37°C in 5% CO2. Following three washes with ice-cold PBS the cells were solubilized in ice-cold RIPA buffer (10 mM Tris-HCl, pH 7.4, 0.15 M NaCl, 1% Triton X-100, 0.1% SDS, 0.65 mM MgSO4, 1 mM CaCl2, 0.5% deoxycholate) containing protease inhibitors (10 turbidity units aprotinin/ml, 0.25 mg DNase I/ml, 0.1 mM Pefabloc, 1 μg Leupeptin/ml, and 1 μM Pepstatin) for 2 h, at 4°C, using a rocking platform. Lysates were scraped into Eppendorf tubes and centrifuged at 15,100 × g for 30 min at 4°C. Protein concentration of the supernatants was determined using a protein assay kit (Bio-Rad Laboratories, München, Germany). Samples of 50 μg protein were separated by SDS-polyacrylamide gel electrophoresis (PAGE) (10% gel) then electrophoretically transferred to a nitrocellulose membrane (Schleicher and Schuell, Dassel, Germany). Unspecific binding sites on membranes were blocked overnight at 4°C with 10% defatted milk in Tris-buffered saline (TBS; 50 mM Tris HCl, pH 7.4, 200 mM NaCl) containing 0.1% Tween-20, and then the membrane was probed either with 3 μg/ml of monoclonal mouse anti-rat CD44 antibodies 5G8 and 1.1ASML or with polyclonal rabbit anti-RHAMM antibodies R 3.4 diluted 1:500 for 2 h at room temperature. After intensive washing with TBS-Tween the membrane was incubated with peroxidase-conjugated anti-mouse IgG (0.2 μg/ml; Vector Laboratories, Burlingame, CA) or with peroxidase-conjugated anti-rabbit IgG (1:5,000; Amersham, UK), respectively, in TBS-Tween for 1 h at room temperature. After washing the blots were developed using the ECL Western blotting detection system (Amersham, UK) according to manufacturer's instructions. Densitometry of the blots was performed on an enhanced laser densitometer (Ultroscan XL; LKB, Bromma, Sweden) and analyzed using the 1-D GelScan XL 2.1 software (Pharmacia LKB Biotechnology, Uppsala, Sweden).
The hyaluronidase activity in AM lysosomes was determined essentially as described by Tung and associates (26). AM lysates were prepared as described above, but instead of RIPA buffer 50 mM Tris buffer at pH 8 was used containing 0.25 M NaCl, 20 mM benzamidine, 2% Triton X-100 and protease inhibitors. Agarose was dissolved in 0.3 M sodium phosphate buffer, pH 4, by heating in a microwave. Hyaluronan solution (Mr 1.3 × 106) was mixed with the agarose after cooling to 60°C to give a final concentration of 0.8 mg/ml of hyaluronan and 0.8% (wt/vol) of agarose. One hundred μl aliquots of the solution were then transfered per well to a 96-well microplate. After the gel had set at room temperature, samples (1 mg of protein from AM lysates diluted in 0.3 M sodium phosphate buffer, pH 4, and acidified by direct addition of concentrated formic acid in final volume of 100 μl and pH 4) were applied to hyaluronan-agarose gel. After 16 h of incubation at 37°C the samples were removed and 100 μl of 10% (wt/vol) aqueous cetylpyridinium chloride was added to each well. The adsorbance at 600 nm was measured by Titertek MK II automated plate reader (EFLAB, Finland) after incubation at room temperature for 30 min. Hyaluronidase activity of samples was calculated from the standard curve performed with testicular hyaluronidase at pH 7.
AM were cultured in a 12-well plate (1 × 106 cells per well) at 37°C for 24 h in RPMI 1640 serum-free medium containing 1 μg/ml of [3H]hyaluronan. Media and washes were combined and the cell layers were dissolved in 0.3 M NaOH containing 1% SDS for 30 min at room temperature. Media and washings as well as the corresponding cell layers were applied separately to 10 × 130 mm G-50 Sephadex superfine columns (Pharmacia Biotech AB, Uppsala, Sweden) and eluted with 0.05 M Na-acetate buffer, pH 6, containing 0.1 M NaCl. Fractions of 450 μl were collected and subjected to scintillation counting (Pharmacia LKB Wallac). The columns were standardized with [3H]hyaluronan and 3H2O.
The method for liver exclusion from systemic circulation in rats was modified from Engström-Laurent and Hellström (27). Ten IU of heparin was given via the spleen under chloralhydrate-pentobarbital anesthesia. Blood pressure and heart rate were monitored from the right femoral artery. A ligature was applied around the hepatoduodenal ligament containing the portal vein and hepatic artery. Blood samples (200 μl) were collected from v. cava inferior with a heparinized syringe before (0 min) and after ligation with 5-min intervals. Circulating blood volume was substituted by isotonic glucose infusion. Under such conditions stable hemodynamic parameters were recorded during 25–30 min. After last sampling the animals were killed by an anesthetic overdose. Blood samples were centrifuged immediately at 820 × g, 10 min, and hyaluronan content in blood plasma was measured with hyaluronan kit Test-50.
Comparisons among more than two data groups were performed with an analysis of variance (ANOVA) corrected with Bonferroni's post hoc. Comparisons between two data groups were performed with a paired Student's t test, after checking the variances with Fisher's exact test. Statistical significance error was set to 5%. Statistical calculations were performed with StatView 4.12 (Abacus Concepts, Berkeley, CA).
AM cultures at 95–99% purity were established from BALF and lung tissue at days 1, 5, and 14 after intratracheal bleomycin or saline administration. The purified cell cultures exhibited typical macrophage morphology when examined with light microscope. Cell cultures obtained at different periods of time from bleomycin-treated or control rats did not differ morphologically. Staining of the cells with ED1 antibodies indicated their macrophage origin (data not shown).
Specific hyaluronan binding sites were quantified on BALF- and tissue-AM tissue isolated from bleomycin-treated and control rats at days 1, 5, or 14 (Figure 2). The cells were incubated with various concentrations of [3H]hyaluronan at 4°C to avoid internalization. The binding of radioactively labeled hyaluronan was inhibited by about 97% in the presence of a 200-fold excess of unlabeled hyaluronan. The hyaluronan binding capacity of AM derived from BALF and lung tissue was almost the same when established from the same animal. The ability of AM from bleomycin-treated rats at days 1 and 5 to bind [3H]hyaluronan was about 8- and 15-fold less, respectively, compared with control counterparts (Figures 2a and 2b). The hyaluronan binding sites in both BALF- and tissue-AM from control rats at day 5 were shown to be saturated with about 3,000 [3H]hyaluronan molecules bound per cell, respectively, with a K d of 0.2 × 10−9 M. The BALF- and tissue-AM from bleomycin-treated rats at day 5 could bind only about 200 [3H]hyaluronan molecules per cell with a K d of 0.3 × 10−9. At day 14 the ability of AM established from bleomycin-treated rats to bind hyaluronan reached almost the control level (Figure 2c). Additional experiments were performed with non-purified cell suspensions obtained from rats at day 5 after bleomycin or saline administration to determine other possible hyaluronan-binding cells. No difference was seen compared with purified AM cultures (data not shown). Binding data was confirmed by incubation of the AM with fluorescein-labeled hyaluronan under the same experimental conditions (Figure 3) with the same cell density on all coverslips. Considerably fewer AM from bleomycin-treated rats possessed hyaluronan binding compared to the control counterparts.
Fig. 2. Specific cell surface hyaluronan receptors on the BALF- and tissue-AM. 2 × 105 BALF-AM (closed circles; open circles) and tissue-AM (closed squares; open squares) from bleomycin-treated (closed circles; closed squares) and control rats (open circles; open squares) were incubated with increasing concentrations of [3H]hyaluronan for 16 h at 4°C. Data represent the mean of duplicates ± variation from a representative experiment after subtraction of unspecific binding at days 1 (a), 5 (b) and 14 (c) after treatment.
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Fig. 3. Hyaluronan binding capacity of BALF- and tissue-AM at day 5. 2 × 105 BALF-AM (a, b) and tissue-AM (c, d) from control (a, c) and bleomycin-treated rats (b, d) were incubated with fluorescein-labeled hyaluronan (100 μg/ml, Mr 1.3 × 106) in RPMI 1640 for 16 h at 4°C. Cell densities on coverslips were quantified with a light microscope and bound hyaluronan with fluorescence optics (Nikon, Japan). Bars = 40 μm.
[More] [Minimize]In order to explore whether the hyaluronan receptors CD44 or RHAMM are responsible for the hyaluronan binding activity detected on AM their expression on these cells was investigated. Using monoclonal anti-rat CD44 antibody 5G8 (recognizes all CD44 except rare cases when exon 15 is missing) as a probe in Western blot analysis, a 30% increase in the total amount of the expression of CD44 molecules on BALF-AM from bleomycin-treated rats at day 5 compared with that of control rats was detected (Figure 4). The amount of CD44 molecules on tissue-AM from bleomycin-treated rats was about 50% higher compared with the control rats (data not shown). The CD44 variants expressed on AM from control and bleomycin-treated rats revealed molecular masses of 74–89 kD and 67–79 kD, respectively. No expression of CD44v6 or RHAMM could be detected on AM by immunoblotting using 1.1ASML and R 3.4 antibodies, respectively.
Fig. 4. CD44 molecules on the BALF-AM at day 5. BALF-AM from three control (lines 1–3) and three bleomycin-treated rats (lines 4–6) were incubated for 24 h in serum-free RPMI 1640 medium. (a) Cell lysates (50 μg of protein) were subjected to SDS-PAGE, transferred to a nitrocellulose membrane and then probed with 5G8 anti-rat CD44 antibody. (b) Densitometric analysis of an immunoblot showing the relative amounts of CD44 molecules (65–90 kD band).
[More] [Minimize]AM have been described to internalize hyaluronan for degradation (4). Therefore we studied lysosomal hyaluronidase activities in BALF- and tissue-AM from bleomycin-treated and control rats at days 1, 5, and 14 (Table 1). Hyaluronidase activity was correlated to the protein content of AM lysates because no marked changes in total protein synthesis have been shown to occur in response to bleomycin treatment (28). Hyaluronidase levels in AM established from bleomycin-treated rats at days 1 and 5 were slightly but insignificantly higher than those from control counterparts. Furthermore, no significant differences in BALF- and tissue-AM hyaluronidase levels were seen at any time point either in bleomycin-treated or control rat groups. Additional experiments were performed with AM lysates prepared with detergent-free buffers, but no difference was seen compared with detergent-containing lysates (data not shown).
| Hyaluronidase activity (units/mg protein) | ||||||
|---|---|---|---|---|---|---|
| Treatment | BALF-AM | Tissue-AM | ||||
| Day 1 | Control | 1.20 ± 0.02 | 1.16 ± 0.02 | |||
| Bleomycin | 1.25 ± 0.01 | 1.22 ± 0.04 | ||||
| Day 5 | Control | 1.21 ± 0.02 | 1.16 ± 0.00 | |||
| Bleomycin | 1.29 ± 0.01 | 1.26 ± 0.03 | ||||
| Day 14 | Control | 1.29 ± 0.01 | 1.27 ± 0.06 | |||
| Bleomycin | 1.25 ± 0.02 | 1.27 ± 0.02 | ||||
Our present and previous findings that AM from bleomycin-treated and control rats express different amounts of functionally active hyaluronan binding sites (18), together with the observation that CD44 participates in the uptake and degradation of hyaluronan (4), prompted us to investigate whether AM from bleomycin-treated and control rats possess different capacities to internalize hyaluronan and degrade it. We found that BALF-AM isolated from bleomycin-treated rats at day 1 were able to internalize 2.3-fold less and degrade 3.0-fold less [3H]hyaluronan, respectively, during a 24-h incubation at 37°C compared with their control counterparts (Figure 5a). BALF-AM from bleomycin-treated rats at day 5 were able to internalize 1.5-fold less and degrade 1.4-fold less [3H]hyaluronan, respectively (Figure 5a). Tissue-AM followed the same pattern and no significant difference was found compared with the corresponding BALF-AM (Figure 5b). At day 14 the ability of AM from bleomycin-treated rats to internalize and degrade hyaluronan was markedly improved, but did not reach the control values yet.
Fig. 5. Internalization and degradation of hyaluronan by BALF- and tissue-AM. 1 × 106 BALF-AM (a) and tissue-AM (b) were cultured for 16 h at 37°C in the presence of 1 μg/ml [3H]hyaluronan. Media combined with the corresponding washes and cell lysates were fractionated separately on G-50 Sephadex superfine column, which was standardized with [3H]hyaluronan and 3H2O. The radioactivity collected in 3H2O defined position was combined and regarded as a total degraded hyaluronan (diagonal lined bars). The cell-associated radioactivity eluted at V0 together with the total degraded hyaluronan was regarded as having been internalized hyaluronan (open bars). Data represent the mean ± SEM from five different animals. *P < 0.05 compared with the group of control animals.
[More] [Minimize]In order to determine whether bleomycin treatment affected circulating hyaluronan uptake by the liver endothelial cells, the amounts of hyaluronan in blood plasma obtained from 6 bleomycin-treated and 6 control rats at day 5 were measured (Figure 6). A total of 177 ± 29 ng hyaluronan per ml blood plasma was observed in control rats and 146 ± 26 ng/ml in bleomycin-treated rats at 0 min, which are similar to the amounts previously observed in untreated rats (reference value 48–260 ng/ml; 29). The whole body lymph node filtration of hyaluronan was indirectly measured by liver exclusion from systemic circulation to block hyaluronan uptake by liver endothelial cells. Rapid accumulation of hyaluronan in circulation was seen in both animal groups after liver exclusion, resulting in about 6-fold elevation at 20 min. Based on the accumulation curve, the average release of hyaluronan to circulation was calculated to be 42.5 ng hyaluronan/ml blood plasma/min in control and 37.5 ng hyaluronan/ml blood plasma/min in bleomycin-treated rats.
Fig. 6. Circulating hyaluronan after liver exclusion at day 5. 200 μl blood samples were collected from bleomycin- (line with closed circles) and saline-treated rats (line with open circles) with 5-min intervals after portal vein ligation. Hyaluronan content in blood plasma was measured in duplicates using hyaluronan kit. Each point represents the mean ± SEM of six different animals.
[More] [Minimize]Accumulation of hyaluronan in BALF and lung interstitium in patients has been described in several lung disorders, such as adult respiratory distress syndrome (30), sarcoidosis (31), and idiopathic pulmonary fibrosis (32). Bleomycin-induced lung injury is a well-established animal model for studies of mechanisms responsible for the development of pulmonary fibrosis. A transient accumulation of hyaluronan in lung interstitium after single intratracheal instillation of bleomycin occurs in early phase of this disorder, peaking at day 5, which is paralleled with water accumulation and inflammatory cell influx (19, 20, 33). Many cell types are potential participants in this process; however, AM play an essential role in acute tissue inflammatory response. There is a concensus that in response to bleomycin treatment the AM become activated and release factors, which stimulate hyaluronan production in lung fibroblasts. Not much is known about the local turnover and the lymph node filtration of hyaluronan which are supposed to be the main mechanisms of hyaluronan breakdown and degradation in peripheral tissues. Totally, about 40–45 μg of hyaluronan is found in lungs of a normal rat (29). In the early phase of fibrotic lung injury the total amount of hyaluronan in the lungs varies between 75 and 105 μg (33). In such a pathological condition with hyaluronan overproduction, the degradatory system fails to maintain the balance. Studies performed in normal hamster AM (4) and human lung fibroblasts (17) demonstrated that both cell types express hyaluronan receptors and are able to internalize and degrade hyaluronan. However, AM possess more hyaluronan binding sites on their cell surface, and degrade hyaluronan much faster than fibroblasts, and may hence account for a major share of the hyaluronan degradation in lungs. To our knowledge, no studies have been performed to evaluate the capacity of AM to degrade hyaluronan in a diseased lung.
To further understand the mechanism of local hyaluronan turnover during the early phase of bleomycin-induced fibrotic lung injury, we isolated and purified BALF- and tissue-AM from bleomycin-treated as well as control rat lungs and studied their ability to internalize and degrade hyaluronan. Tissue-AM have been known also as interstitial macrophages (34, 35), but based on recent studies they are morphologically and histochemically identical to BALF-AM (22). In this context we wanted to explore whether these two macrophage populations differ in their ability to regulate hyaluronan turnover. The genuine interstitial macrophages can be obtained by enzymatical digestion of lung tissue, but as they constitute only 2–5% of the total lung cell population, and exhibit lower functional activity (36, 37), they were excluded from this study.
The mechanisms of hyaluronan turnover and catabolism in normal tissues are well described. First, hyaluronan has to be specifically bound to the surface of cells with hyaluronan degradatory capacity. Specific hyaluronan receptors have been described on normal AM which were defined as CD44 molecules (4, 5, 38). CD44 on the surface of inflammatory macrophages undergoes posttranslational modifications including N-linked glycosylation and phosphorylation, which is not seen in resident cells (39). There is also evidence that conformational changes of CD44 may be required for hyaluronan binding (40). In this context, our finding that increased amounts of lower molecular mass CD44 molecules could be detected on AM from bleomycin-treated rats compared with control rats (Figure 4), with a concomitant decrease in hyaluronan binding activity (Figure 2) is of particular interest. This observation suggests that modifications of the CD44 molecule, which are needed for hyaluronan binding, do not occur in diseased state. The lack of blocking antibodies against rat CD44 makes it difficult to determine whether all hyaluronan binding activity on AM is due to CD44 or whether other, yet undefinable hyaluronan binding proteins are involved.
Second, hyaluronan has to be internalized and transported to lysosomes. Our results indicate a decrease of hyaluronan internalization already at day 1 after induction of lung injury. The capacity of internalization improved gradually to day 14, which also can be explained by maturation of a new AM cell generation. Strong correlation was found between the number of specific hyaluronan binding sites (Figure 2) and hyaluronan internalization capacity (Figure 5) of AM which indicates the primary role of hyaluronan ligation to specific receptors as an initial step for local hyaluronan degradation. Moderate decrease of hyaluronan internalization capacity was seen in control rats at day 5 as well, indicating that tracheal cannulation together with saline instillation is sufficient to induce minor lung injury and affect the hyaluronan balance in lung interstitium.
Third, hyaluronan is subjected to enzymatic digestion in lysosomes, where it is digested to oligosaccharides by hyaluronidase, and then to monosaccharides by β-D-glucuronidase and N-acetyl-β-D-hexosaminidase (11). The results of this study show that only about 50% of internalized hyaluronan is degraded at any time point after intratracheal treatment both in injured and normal lungs (Figure 5). The finding that lysosomal hyaluronidase activities in AM established from bleomycin-treated and control rats did not differ significantly (Table 1) indicates that the intracellular mechanism for hyaluronan degradation remains unaffected in lung injury. Interestingly, despite a 15-fold decrease of hyaluronan binding capacity in bleomycin-treated rats at day 5, the ability of AM to internalize and degrade hyaluronan was decreased only by 35%. This may indicate that the few functional hyaluronan receptors on the AM surface obtained from bleomycin-treated rats are more efficient in their internalizing capacity than those on AM from control animals, or that other phagocytotic mechanisms which do not require specific binding of hyaluronan to receptors are involved.
Another pathway for hyaluronan turnover is catabolism or fragmentation during filtration in lymph nodes. Increased circulating hyaluronan levels are found in several pathological conditions with hyaluronan overproduction or decreased clearance by liver endothelial cells (for review, see reference 41). Our finding that circulating hyaluronan levels in rats with bleomycin-induced lung injury are not elevated (Figure 6; 0 min) indicate that hyaluronan uptake by liver endothelial cells is not affected by bleomycin treatment. We detected also a slightly decreased release of hyaluronan to the circulation in bleomycin-treated rats at day 5, when the hyaluronan levels are known to peak in lung interstitium (19). Accumulation of hyaluronan in lung interstitium may alter the osmotic pressure and thereby decrease transinterstitial fluid flux, which has been shown to determine the lymphatic output of hyaluronan (42).
The findings of the present study further support the theory that accumulation of hyaluronan in lung interstitium during the early phase of fibrotic lung injury is due to both an overproduction of hyaluronan by lung fibroblasts and to an impairment of the function of hyaluronan receptors on AM.
The writers thank Prof. T. C. Laurent and Dr. O. Nettelbladt for support and constructive criticism of this work. This project was supported in part by grants from The Swedish Cancer Foundation, The Swedish Heart Lung Foundation, Gustaf V:s 80-års fond, and The Swedish Medical Research Council (03X-4).
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