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Published ahead of print on June 12, 2003, doi:10.1165/rcmb.2003-0078OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 29, pp. 743-749, 2003
© 2003 American Thoracic Society
DOI: 10.1165/rcmb.2003-0078OC

Pulmonary and Activation-Regulated Chemokine Stimulates Collagen Production in Lung Fibroblasts

Sergei P. Atamas*, Irina G. Luzina*, Jung Choi, Natalya Tsymbalyuk, Nicholas H. Carbonetti, Ishwar S. Singh, Maria Trojanowska, Sergio A. Jimenez and Barbara White

Departments of Medicine and of Microbiology and Immunology, University of Maryland School of Medicine; and Research Service, Veterans Affairs Maryland Health Care System, Baltimore, Maryland; Division of Rheumatology and Immunology, Medical University of South Carolina, Charleston, South Carolina; and Department of Medicine, Jefferson Medical College, Thomas Jefferson University, Philadelphia, Pennsylvania

Address correspondence to: Sergei P. Atamas, M.D., Ph.D., Baltimore VA Medical Center, Research Service (151), Room 3C-125, 10 North Greene Street, Baltimore, MD 21201. E-mail: satamas{at}umaryland.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Levels of pulmonary and activation-regulated chemokine (PARC) mRNA and protein are increased in the lungs of patients with pulmonary fibrosis. The purpose of this study was to establish whether PARC could be directly involved in development of pulmonary fibrosis by stimulating collagen production in lung fibroblasts. Exposure to PARC increased production of collagen mRNA and protein by 3- to 4-fold in normal adult lung and dermal fibroblast cells. Collagen mRNA transiently increased after 3–6 h of activation with PARC, with an increase in collagen protein detected after 24 h of activation. At the same time, PARC had less pronounced effect on fibroblast proliferation, not exceeding 50% increase over control nonstimulated cells. PARC intracellular signaling led to activation of ERK1/2, but not p38, in fibroblasts; pharmacologic inhibition of ERK, but not p38, also blocked PARC's effect on collagen production. Inhibition experiments with pertussis toxin suggested that PARC receptor is G protein–coupled. Thus, PARC is a member of the CC chemokine family that acts directly as a profibrotic factor.

Abbreviations: bronchoalveolar lavage, BAL • enzyme-linked immunosorbent assay, ELISA • extracellular signal-regulated kinase, ERK • interleukin, IL • monocyte chemotactic protein, MCP • macrophage inflammatory protein, MIP • pulmonary and activation-regulated chemokine, PARC • polymerase chain reaction, PCR • pertussis toxin, PT • recombinant human, rh • transforming growth factor-ß1, TGF-ß1


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Increased levels of pulmonary and activation-regulated chemokine (PARC) mRNA and protein have been previously found in patients with scleroderma lung disease, a condition associated with lung fibrosis (1). In hypersensitivity pneumonitis, which frequently leads to lung fibrosis, and in idiopathic pulmonary fibrosis, levels of PARC are also increased (2). An increase in PARC mRNA levels was found in the lung tissue in a monkey model of asthma (3). This finding raises the possibility that PARC expression may stimulate the subepithelial fibrosis that underlies airway remodeling in asthma. Also, PARC mRNA is increased in the lungs of patients with sarcoidosis, a condition in which fibroblasts contribute to collagen deposition and subsequent lung fibrosis (4).

This chemokine is also known as macrophage inflammatory protein (MIP)-4, alternative macrophage activation-associated CC chemokine (AMAC)-1, dendritic cell chemokine (DC-CK)-1, CC chemokine ligand (CCL)-18, small secreted cytokine A (SCYA)-18, and chemokine ß7 (Ckß7). PARC is a chemotactic factor for T cells that is constitutively expressed at high levels in the lungs (5), particularly by alveolar macrophages (2, 6). It can also be produced, although to a lesser degree, by peripheral blood monocytes, tissue macrophages, and dendritic cells (2, 6–9). Resting and activated fibroblasts do not produce PARC (9). PARC is chemotactic for both naive and activated CD4+ and CD8+ T cells, but not granulocytes or monocytes (5, 7). Based on the observed association between PARC expression and lung fibrosis, the hypothesis of this work was that PARC, besides attracting T cells, could also stimulate collagen production and/or fibroblast proliferation.

We show here that PARC activates collagen production in cultured human lung and dermal fibroblasts derived from normal adult healthy donors. An increase in type I mRNA and protein collagen levels has been observed in response to stimulation of cultured fibroblasts with PARC. Preceding the increase in collagen production, activation of the extracellular signal-regulated kinse (ERK) pathway through a G protein–coupled receptor takes place. These observations, together with previous reports of increased PARC levels in association with lung fibrosis, suggest that this chemokine is a direct profibrotic factor, particularly in the lung tissue.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Recombinant Human Cytokines
Recombinant human (rh) PARC and rhIL-4 were purchased from R&D Systems (Minneapolis, MN). Carrier-free rhPARC was purchased from Cell Sciences (Norwood, MA). Neutralizing anti-human PARC antibody was purchased from R&D Systems.

Fibroblast Cell Lines
Four normal human lung fibroblast lines (LF1-LF4) derived from primary lung explants from adult donors were purchased from BioWhittaker (Walkersville, MD). Three normal adult human dermal fibroblast lines (DF1–DF3) were previously established in the laboratory from primary dermal explants, as described (10). Fibroblast lines were maintained in T75 culture flasks in humidified atmosphere of 5% CO2 at 37°C in high-serum tissue culture medium, which was Dulbecco's modified Eagle's medium supplemented with 10% bovine calf serum, 2 mM glutamine, 2 mM sodium pyruvate, and 50 mg/liter gentamicin (all from Life Technologies, Grand Island, NY). Before experiments, cell cultures were preincubated for 24 h in similar conditions, except that low-serum (0.5% dialyzed bovine calf serum with no TGF-ß detectable by enzyme-linked immunosorbent assay [ELISA] as described below) medium was used, supplemented in addition to the mentioned reagents with 0.28 mM ascorbic acid and 0.2 mM ß-aminopropionitrile (Sigma, St. Louis, MO). Cell culture medium for all experiments was the same low-serum medium. In all experiments, fibroblast cell lines were tested in passages three to seven.

Real-Time Polymerase Chain Reaction for Collagen mRNA
Total RNA was purified from fibroblast monolayers using Trizol (Gibco Invitrogen, Carlsbad, CA), as described (11). Collagen mRNA was measured by real-time polymerase chain reaction (PCR) (LightCycler; Roche, Indianapolis, IN). The primers and hybridization probes for {alpha}2(I) collagen mRNA and reference sequence 18S rRNA were prepared by TIB Molbiol (Adelphia, NJ). The primers for {alpha}2(I) collagen were: forward, 5'-GAT GGT GAA GAT GGT CCC ACA GG-3', and reverse, 5'-GGT CGT CCG GGT TTT CCA GGG T-3'. The hybridization probes for {alpha}2(I) collagen were labeled with fluorescein at the 3'-terminus (3FL) of one probe and with LightCycler Red (5LC) at the 5'-terminus of the other probe. The probes were: 3FL 5'-TTC CAA GGA CCT GCT GGT GAG CCT-3'FL and 5LC 5'-TGA ACC TGG TCA AAC TGG TCC TGC AG-3'. The PCR reaction mixture included 5 mM MgCl2, 0.5 µM primers, 0.2 µM probes, and the recommended components of the FastStart DNA Master Hybridization Probes hot start reaction mix (Roche). The following PCR conditions were used for {alpha}2(I) collagen: activation at 95°C for 10 min, followed by 35 cycles of denaturation at 95°C for 3 s, primer annealing at 61°C for 5 s, fluorescence readout (Fl2/Fl1) at 68°C, and extension at 72°C for 15 s. Primers, PCR protocol, and product quantification for 18S rRNA were exactly as reported previously (12). The amount of {alpha}2(I) collagen mRNA was expressed relative to the amount of 18S rRNA in the same sample. Actinomycin D (transcription inhibitor) was purchased from Sigma. Cultures were tested in duplicate in three independent experiments for reach cell line, and data were analyzed using one-way ANOVA with post hoc (Scheffe) testing utilizing Statistica software (StatSoft, Tulsa, OK).

Collagen Production Assays
Production of collagen was assessed using metabolic labeling with 14C-proline and Western blotting with anti-collagen type I antibody. For metabolic labeling of collagen, fibroblasts were plated at 2 x 105 cells/well in 6-well plates (Costar, Cambridge, MA), in duplicates, incubated overnight in 3 ml/well of high serum medium, and then for 24 h in low-serum medium. After that, the culture medium was replaced with 1 ml/well of fresh low serum medium with or without added test substances and containing 14C-proline at 1 µCi/ml (Amersham Pharmacia Biotech). After incubation for the desired periods of time, the cell culture supernatants were collected, rapidly frozen in liquid nitrogen and freeze-dried at -70°C. The pellets were dissolved in 100 µl of reducing Laemmli buffer per 1 ml of the cell culture supernatant, and the samples were electrophoretically separated in 7.5% acrylamide gels. Alternatively, samples were concentrated 10-fold by filter centrifugation on 30K Ultrafree-MC filters (Millipore, Bedford, MA). Fluorographic images were developed using EN[3H]ANCE autoragiography enhancer (NEN, Boston, MA). Gel images were acquired using Storm densitometer (Molecular Dynamics, Sunnyvale, CA), and the densities of the bands were analyzed with ImageQuant software (Molecular Dynamics). Dependence of collagen production by fibroblast cultures on the dose of PARC and duration of stimulation were evaluated using one-way ANOVA with post hoc testing.

Western blotting assays for collagen were performed using rabbit affinity purified anti-collagen type I antibody (Rockland, Gilbertsville, PA). Before electrophoresis, samples were reduced and denatured by boiling in Laemmli buffer containing ß-mercaptoethanol. Human purified collagen type I (Southern Biotech, Birmingham, AL) was used as a positive control in these assays.

The identity of collagen bands was confirmed by sensitivity to pepsin and collagenase (both from Sigma) digestion. For pepsin digestion, 2 µl of 1M acetic acid were added to 40 µl of concentrated fibroblast culture supernatant, followed by 1 µl of pepsin stock, to achieve final concentration of pepsin as indicated in the text. After 15 min of digestion at room temperature, reaction was stopped with 4 µl of 1 M Tris base and 40 µl of reducing Laemmli buffer. For collagenase digestion, 1 µl of 4 U/ml bacterial collagenase and 1 µl of protease inhibitor cocktail (Sigma) were added to 40 µl of sample and digestion performed at room temperature for 30 min. The bacterial collagenase was highly purified and had minimal clostripain and neutral protease activity, as tested by the manufacturer, according to the product information sheet.

Cell Proliferation Assays
For CellTiter AQueous 96 Non-Radioactive Cell Proliferation Assay (Promega, Madison, WI), fibroblasts were plated in low-serum medium at 2 x 103 cells/well in 96-well flat-bottom tissue culture plates (Costar) in 0.2-ml cultures and stimulated with increasing concentrations of rhPARC in quadruplicates. Low-serum tissue culture medium alone was the negative control. The CellTiter proliferation assay was performed according to the manufacturer's instructions, after the fibroblasts were incubated with test substances for 3–8 d. Data were expressed as mean OD490 ± SD of quadriplicate cultures and analyzed using one-way ANOVA with post hoc testing.

Immunoblotting for Phosphorylation of EKR1/2 and p38
Fibroblasts were plated in 6-well plates (Costar) at 2 x 105 cells/well in 3-ml cultures. After incubation with PARC for 15 min, fibroblast cultures were washed with ice-cold phosphate-buffered saline containing 100 µM Na3VO4. Then, fibroblasts were lysed with 250 µl of Laemmli sample buffer. Electrophoretic separation of cell lysates was done in 7.5% acrylamide gels, and bands were transferred onto Immobilon NC membranes (Millipore, Bedford, MA). Membranes were probed with specific primary antibodies at 1/200 dilution, then secondary goat anti-mouse IgG-HRP conjugate (Upstate, Lake Placid, NY), and visualized with an ECL detection system (Pierce, Rockford, IL) that was used according to the manufacturer's directions. Anti-ERK, anti-P38, anti–phospho-ERK1/2 and anti–phospho-p38 mAb were purchased from Upstate. Gel images were collected using a Storm densitometer and band densities analyzed with ImageQuant software (Molecular Dynamics). ERK inhibitor PD98059 and p38 inhibitor SB203580 purchased from Upstate were > 98% chromatographically pure and quality control tested by the supplier, and confirmed to selectively inhibit their target enzymes. Cell viability in the presence of inhibitors was determined using Trypan Blue exclusion assays.

Inhibition of Receptor Signaling with Bordetella Pertussis Toxin
Wild-type pertussis toxin (PT) and inactive mutant PT (PT9K/129G) were purified from Bordetella pertussis W28 culture supernatant by fetuin affinity chromatography as previously described (13). Both wild-type and inactive mutant PT were added to fibroblast cell cultures in final concentration of 10 ng/ml, fibroblasts were stimulated with rhPARC, and ERK1/2 phosphorylation tested by Western blotting after 15 min of activation with PARC.

Transforming Growth Factor-ß1 ELISA Assays
ELISA kits for transforming growth factor (TGF)-ß1 were purchased from R&D Systems (Minneapolis, MN), and fibroblast culture supernatants and whole cell lysates after 3, 6, 12, 24, 48, and 72 h activation with rhPARC were assayed in duplicates for total TGF-ß1, according to the manufacturer's instructions. Low-serum cell culture medium containing 0.5% dialysed fetal bovine serum had no detectable TGF-ß1 and was used as a negative control in these assays.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Production of Type I Collagen Protein Is Increased in Response to PARC
To test effects of rhPARC on collagen protein production by lung and dermal fibroblast cell lines, fibroblast cultures were incubated for 48 h with or without 30 ng/ml and 300 ng/ml rhPARC in the low-serum cell culture medium containing 14C-proline for metabolic labeling of collagen. Fibroblast culture supernatants contained two 14C-proline containing bands in gel electrophoresis under reducing conditions (Figures 1A and 1B). Digestion with chromatographically pure bacterial collagenase in the presence of protease inhibitors eliminated the bands completely (Figure 1A). To confirm the identity of these bands, we conducted Western blotting experiments in reducing conditions of fibroblast culture supernatants using an anti-collagen type I antibody, that recognizes {alpha}1(I) collagen but not other collagen chains (14). Digestion with pepsin caused expected decrease in the apparent molecular weight of the immunoreactive collagen from ~ 175 kD to ~ 125 kD (14) (Figure 1C).



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  		Figure 1. Collagen production after activation of fibroblast cell cultures with recombinant human PARC for 48 h. (A) Collagen was metabolically labeled with 14C-proline in LF1 cells, culture supernatant separated in PAGE under reducing conditions, and collagen chains visualized by fluorographically enhanced autoradiography. Equal loading was ensured by adjusting total protein in the loaded sample. Samples were loaded as follows: lane 1, control nonstimulated fibroblast supernatant; lane 2, supernatant from fibroblast culture activated with 300 ng/ml rhPARC; and lane 3, sample 2 digested with collagenase. The combined density of procollagen bands in lane 2 is 3.4-fold higher than in lane 1, after adjustment to the local background. (B) Collagen was metabolically labeled with 14C-proline in LF2 cells, fibroblast culture supernatants subjected to PAGE under reducing conditions, and bands visualized as outlined above in A. Samples were loaded as follows: lane 1, control nonstimulated fibroblast culture supernatant; lane 2, supernatant from fibroblasts activated with 30 ng/ml rhPARC; lane 3, supernatant from fibroblasts activated with 300 ng/ml rhPARC; and lane 4, supernatant from fibroblasts activated with 10 ng/ml rhIL-4. The combined density of procollagen bands in lanes 2, 3, and 4 are 2.7-, 4.4-, and 2.9-fold higher, respectively, than in lane 1, after adjustment to the local background. (C) Western blotting of LF2 cell culture supernatants for collagen {alpha}1(I). Samples were loaded in the following order: lane 1, human type I collagen (Southern Biotech); lane 2, control nonstimulated fibroblast supernatant; lane 3, supernatant from fibroblasts activated with 300 ng/ml rhPARC; lanes 4–6, sample 3 digested with 125 µg/ml, 25 µg/ml, and 2.5 µg/ml pepsin, respectively. The combined density of procollagen bands in lane 3 is 4.2-fold higher than in lane 2, after adjustment to the local background. (D) Western blotting of LF4 cell culture supernatant for collagen {alpha}1(I). Samples were loaded as follows: lane 1, control nonstimulated fibroblast supernatant; lane 2, supernatant from fibroblasts activated with 100 ng/ml rhPARC; lane 3, same as sample 2 incubated with 100 µg/ml neutralizing anti-PARC antibody. The combined density of procollagen bands in lanes 2 and 3 are 3.6- and 1.4-fold higher, respectively, than in lane 1, after adjustment to the local background.

 
Exposure to rhPARC from both sources (R&D Systems and Cell Sciences) for 48 h increased fibroblast production of collagen more than 3-fold over control in five out of seven tested fibroblast lines (Figure 1). The stimulating effect of rhPARC on collagen production was inhibited by adding 100 µg/ml of neutralizing anti-PARC antibody to the cell cultures (Figure 1D). Three lung fibroblast lines and two dermal lines, each tested in at least three experiments, all responded to rhPARC stimulation, with a dose of 300 ng/ml consistently stimulating collagen production. The average increase in the density of collagen bands after 48 h of stimulation with 300 ng/ml rhPARC was 3.4 ± 0.9 fold (P < 0.01, Student's t test, compared with control unstimulated cultures). One lung fibroblast line and one dermal fibroblast line were consistently nonresponsive to rhPARC stimulation up to 72 h and rhPARC concentrations up to 1,000 ng/ml.

Further experiments defined the dose–response (Figure 2A) and the kinetics of PARC's effect (Figure 2B) on collagen production in LF1 and LF4 lines, with a total of five experiments done with similar results. One-way ANOVA analyses with post hoc testing revealed a significant increase in collagen production in response to 30 ng/ml (P < 0.05) and 300 ng/ml (P < 0.01) rhPARC, with increases also seen at 3 ng/ml (P < 0.12) and 10 ng/ml (P < 0.1) rhPARC. There was no difference in collagen production between control cultures and those exposed to PARC for 3, 6, and 12 h of activation, with increase in collagen production not exceeding 1.2- ± 0.2-fold over control nontreated cells (P > 0.05). A significant increase in collagen production (P < 0.05) was observed after 24, 48, and 72 h of activation with rhPARC (Figure 2B).



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  		Figure 2. Increase in production of collagen depends on the dose of PARC and time of activation. (A and B) Western blotting of LF4 cell culture supernatants for collagen {alpha}1(I) (upper part of each panel) and densitometry of the combined pro-{alpha}1(I) collagen bands adjusted to local background (lower part of each panel). (A) Fibroblasts were activated for 48 h with increasing concentration of rhPARC from 1 ng/ml to 300 ng/ml. (B) Fibroblast cultures were incubated for 24, 48, and 72 h without (Ctrl) or with 300 ng/ml rhPARC (PARC). The increase in densities of the collagen bands relative to the control culture was 4.4 at 24 h, 3.2 at 48 h, and 1.3 at 72 h, although densities of both pro-{alpha}1(I) collagen bands in the PARC-treated sample at 72 h of activation were saturated and underrepresented the true increase in collagen levels. Densitometry of the same bands measured at a shorter, nonsaturating exposure, showed a 3.4-fold increase in collagen production by rhPARC-stimulated fibroblasts at 72 h.

 
Collagen {alpha}2(I) mRNA Is Increased in Response to PARC
Experiments were done to test whether an increase in steady-state collagen mRNA levels might be a mechanism of PARC stimulation of collagen protein production. Fibroblast lines LF1 and LF2 were incubated in the low-serum cell culture medium with and without 300 ng/ml rhPARC for 0 min, 90 min, 3 h, 6 h, and 24 h. Total mRNA was purified and collagen mRNA levels were tested in a semiquantitative RT-PCR with levels of collagen mRNA determined relative to 18S rRNA levels. The steady state mRNA levels for {alpha}2(I) collagen chain were transiently increased between 3 and 6 h from the time of activation (Figure 3). Treatment of cell cultures with 10 µg/ml actinomycin D (inhibitor of transcription) before stimulation with PARC completely abrogated the increase in collagen mRNA (data not shown). Both fibroblast lines were tested each in three separate experiments with similar results.



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  		Figure 3. Real-time PCR of 18S rRNA and collagen mRNA in lung fibroblasts, control and treated with 300 ng/ml of rhPARC. Detection of 18S PCR product was done with SYBR Green (fluorescence 1, Fl1 on the left vertical axis in A) and detection of collagen PCR product was done with specific HybProbes (fluorescence1/fluorescence 2, Fl1/Fl2 on the right vertical axis in A). (A) Equal RNA concentration in control and PARC-treated cells, based on close overlap of corresponding 18S rRNA amplification curves. This panel also shows that after 6 h of incubation in triplicates, amplification of collagen PCR product from fibroblasts treated with rhPARC occurs approximately two cycles earlier than in control samples, indicating approximately 4-fold higher concentration of collagen mRNA in the treated samples. (B) Kinetics of collagen mRNA increase in LF1 (open bars) and LF2 (shaded bars) fibroblasts treated with PARC versus nonstimulated cells incubated for the same periods of time, after normalization to 18S rRNA. Levels of collagen at 3 and 6 h of activation are higher than in control cells (P < 0.05 by one-way ANOVA with post hoc testing), and the differences approach statistical significance at 24 h (P < 0.09).

 
PARC Does Not Stimulate Production of TGF-ß1 from Fibroblasts
To exclude the possibility that collagen production is activated by an increase in autocrine TGF-ß1 production after PARC stimulation, levels of this cytokine were measured by ELISA in supernatants and whole cell lysates from fibroblasts exposed for 3, 6, 12, 24, 48, and 72 h to 300 ng/ml rhPARC. These experiments were repeated in all studied lung and dermal fibroblast lines. Although there was significant variability in baseline TGF-ß1 production between the fibroblast cell lines, the maximal increase of 1.3- ± 0.2-fold in production of TGF-ß1 by fibroblasts (P > 0.05 by Student's t test) was observed in response to PARC activation (not shown).

PARC Has Limited Effect on Fibroblast Proliferation
Effects of PARC on fibroblast proliferation were less pronounced than its effects on collagen production. Fibroblast cultures were incubated with and without rhPARC for 3–8 d. rhPARC was tested in various concentrations ranging from 1–3,000 ng/ml, in quadruplicate cultures. Lung and dermal lines cell lines, LF1, LF2, DF1, and DF2 were tested in proliferation assays, each line in at least two experiments. Although PARC stimulated fibroblast proliferation in these lung and dermal fibroblast lines, the maximum increase over nonstimulated control was 20–25% in lung lines and 45–50% in dermal lines on Days 7 and 8 of incubation (data not shown).

PARC Signals through ERK, but not p38, Pathways
Experiments were done to determine whether rhPARC activated phosphorylation of the MAP kinase pathways. Fibroblasts were activated with rhPARC in low-serum cell culture medium, lysed, and phosphorylation of ERK1/2 and another MAP kinase, p38, was studied by Western blotting. Two independent experiments were done, each of which tested LF1 and LF2 cell lines. Phosphorylation of ERK1/2, but not p38, was increased in lung fibroblasts (Figure 4). ERK pathways appear critical for the effect, because PD98059, a specific inhibitor of ERK activation, also inhibited PARC-activated collagen production in lung fibroblasts, whereas SB203580, a specific inhibitor of p38 activation, had no effect (Figure 5).



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  		Figure 4. Western blotting of whole cell lysates with anti–phospho-ERK1/2 (A), anti–phospho-p38 (B), and ERK2 for loading control (C), after activating lung fibroblasts LF1 with rhPARC for indicated times (min). Phosphorylation of ERK1/2 but not p38 is activated by rhPARC.

 


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  		Figure 5. Collagen production after activation of fibroblast cell cultures LF2 with rhPARC, p38 inhibitor SB203580, and ERK inhibitor PD98059 (A) and densitometric values of the collagen bands in the corresponding lanes (B). Collagen was metabolically labeled with 14C-proline, culture supernatants were separated in PAGE, and procollagen chains were visualized by fluorographically enhanced autoradiography.

 
The PARC Receptor Is G Protein–Coupled
The molecular identity of PARC receptor remains unknown. Experiments were done to confirm that the PARC receptor is G protein–coupled, as are other chemokine receptors (15). LF1 fibroblasts were activated with 300 mg/ml rhPARC for 15 min alone and in the presence of 10 ng/ml Bordetella PT, which inhibits G protein–coupled signaling, or its inactive mutant. Bordetella PT, but not the inactive mutant, inhibited phosphorylation of ERK in LF1 fibroblasts in two independent experiments (Figure 6). These data indicate that the PARC receptor is G protein–coupled, similar to other CC chemokine receptors.



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  		Figure 6. Western blotting of LF1 whole cell lysates with anti–phospho-ERK1/2 (top) and anti-ERK2 for loading control (bottom). Sample 1, control nonstimulated fibroblasts LF1; sample 2, fibroblasts LF1 activated for 15 min with 300 ng/ml rhPARC; sample 3, same as sample 2, plus 10 ng/ml of pertussis toxin; and sample 4, same as sample 2, plus 10 ng/ml of inactive mutant PT. Bordetella PT, but not its inactive mutant, block PARC-stimulated ERK phosphorylation, suggesting that PARC signaling is G protein–coupled.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Pulmonary and activation-regulated chemokine is a CC chemokine that is chemotactic for T cells, but not other cell types (5, 7). As reflected in its name, this chemokine is expressed at high levels in the lungs, particularly by activated lung macrophages (2, 6), although other tissue macrophages and dendritic cells can also secrete PARC. Macrophages activated by interleukin (IL)-4 and TGF-ß, but not interferon-{gamma} or tumor necrosis factor-{alpha} (alternatively activated macrophages, reviewed in Ref. 16) are a source of PARC (6), as reflected in another name for this factor, alternative macrophage activation-associated chemokine.

This work reports that PARC directly stimulates type I collagen production in lung and dermal fibroblasts up to 4-fold over control nonstimulated cells. At the same time, PARC causes only 25–50% increase in fibroblast proliferation. Collagen protein and mRNA both were increased after activation of fibroblasts with PARC (Figures 1–3). This increase in mRNA indicates that an increase in collagene gene transcription or an increase in collagen mRNA stability are potential mechanisms of collagen upregulation by PARC. In addition, it remains possible that some of the increase in collagen protein could be due to an effect of PARC on the intracellular free proline pool.

To investigate possible intracellular signaling events activated by PARC, experiments tested activation of the MAP kinase pathways. Phosphorylation of ERK, but not p38, was increased by PARC in a time-dependent manner (Figure 4). Pharmacologic inhibition of ERK, but not p38, blocked the increased production of collagen by fibroblasts in response to PARC (Figure 5) suggesting that ERK, but not the p38 pathway, is critical for PARC's effect on collagen production. Thus, PARC directly stimulates collagen production in lung and dermal fibroblasts by activating intracellular signaling through the ERK pathway. Although the PARC receptor remains unknown, our data indicate that this receptor is G protein–coupled (Figure 6), as are the receptors for other CC chemokines (15). These observations are in agreement with a previous reports (17, 18) indicating that another CC chemokine, monocyte chemotactic protein (MCP)-1, activates ERK signaling, which is dependent on G protein–coupled signal transduction in monocytes. Unlike PARC, MCP-1 also activated p38 signaling in monocytes, which was independent of G protein–coupled signaling (17). Another CC chemokine, RANTES, signals though ERK pathway in eosinophils (19) and T cells (20). Together, our observations and data from research literature suggest that ERK signaling activated by G protein–coupled receptor signaling may be a common signaling pathway for CC chemokines.

The concentrations of PARC required to achieve a significant effect on collagen production are compatible with those reported for MCP-1 (21), with PARC causing some stimulation of collagen production in concentrations below 30 ng/ml (see Figure 2), and MCP-1 stimulating collagen production in concentrations of 100–400 ng/ml (21). Considering that activated alveolar macrophages are an abundant source of PARC (2, 6) and that lung macrophages are actively involved in lung inflammation leading to fibrosis (1, 2), it is possible that, during lung inflammation, PARC may be present in lung tissue in amounts that stimulate lung fibrosis. Although precise quantitative data on the levels of PARC in the lungs of patients with fibrotic lung conditions are not available, our own observations indicate that PARC levels in the bronchoalveolar lavage (BAL) fluids of patients with scleroderma lung disease can be as high as 12 pg/ml, with average levels of 6 pg/ml. Such levels are comparable with levels of MCP-1 measured in the BAL fluids from the same patients (1). Because airway and alveolar fluids are diluted during the lavage procedure, BAL fluids are likely to under-represent cytokine concentrations present in the lung tissue.

It is a possible that the profibrotic effect of PARC might be mediated by autocrine TGF-ß, as has been suggested for MCP-1 (21). However, no increase in TGF-ß protein was found by ELISA in cell culture supernatants or whole cell lysates of fibroblast lines stimulated for 3–72 h with rhPARC, suggesting that autocrine TGF-ß is unlikely to mediate effects of PARC on collagen production by lung fibroblasts.

The results reported here, in combination with previous reports of increased PARC mRNA and protein in lung fibrosis (1–4), suggest that the CC chemokine PARC is a new member of the group of cytokines that regulate fibroblast activities in a profibrotic manner. Other factors in this group include, but are not limited to, TGF-ß, IL-4, and oncostatin M (reviewed in Ref. 22). Before this report, MCP-1 was the only other CC chemokine known to be capable of increasing collagen production in fibroblasts (21, 23).

IL-4 appears to promote fibrosis through two mechanisms: by directly stimulating collagen production in fibroblasts and by activating other cells to produce profibrotic cytokines. For example, IL-4 and other type 2 cytokines promote alternative activation of lung macrophages that enhance fibrogenesis by providing profibrotic factors, such as TGF-ß, platelet-derived growth factor, and MCP-1 (24). Among cytokines whose production is differentially regulated in classically and alternatively activated macrophages is PARC (6). Interferon-{gamma} inhibits, whereas IL-4, IL-13, and IL-10 induce, PARC production in these cells (6). Thus, alternatively activated macrophages might be important players in the mechanisms of lung fibrosis. The development of pulmonary fibrosis is generally associated with predominant expression of type 2 cytokines in the lungs (reviewed in Ref. 25). This report suggests that type 2 cytokines promote lung fibrosis not only by directly acting on lung fibroblasts, but also by activating lung macrophages through the alternative pathway to increase production of PARC. This work identifies PARC as a profibrotic factor that directly stimulates collagen production and is a potential target for future antifibrotic therapies, especially in pulmonary fibrosis.


   Acknowledgments
 
This work was supported in part by the National Institutes of Health Grants 1R03AR47110 and by a research grant from the Maryland Chapter of Arthritis Foundation (to S.P.A.). The authors thank Dr. Jeffrey D. Hasday (University of Maryland School of Medicine) for valuable discussion of this work.


   Footnotes
 
* These authors contributed equally to this work. Back

Received in original form March 13, 2003

Received in final form June 3, 2003


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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