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Inhibitory Effects of Pirfenidone on Dendritic Cells and Lung Allograft Rejection

Bizargity, Peyman1; Liu, Kaifeng1; Wang, Liqing2; Hancock, Wayne W.2; Visner, Gary A.1,3

doi: 10.1097/TP.0b013e3182584879
Basic and Experimental Research

Background Pirfenidone (PFD) is an antifibrotic agent with beneficial effects on proinflammatory disorders. In this study, we further investigated PFD and long-acting form, “deuterated PFD,” immune-modulating properties by evaluating their effects on mouse dendritic cells (DCs).

Methods The effects of PFD on DCs were examined in vivo using an orthotopic mouse lung transplant model and in vitro using isolated bone marrow–derived DCs in response to lipopolysaccharide and allogeneic stimulation.

Results In mouse lung transplants, PFD and deuterated PFD treatment improved allograft lung function based on peak airway pressure, less infiltrates/consolidation on micro–computed tomography scan imaging, and reduced lung rejection/injury. DC activation from lung allografts was suppressed with PFD, and there seemed to be a greater effect of PFD on CD11c+CD11bCD103+ lung DCs. In addition, PFD reduced the expression of several proinflammatory cytokines/chemokines from lung allografts. In vitro, DCs treated with PFD showed decreased expression of major histocompatibility complex class II and costimulatory molecules and the capacity of these DCs to stimulate T-cell activation was impaired, although antigen uptake was preserved. PFD directly inhibited the release of inflammatory cytokines from isolated DCs, was associated with a reduction of stress protein kinases, and attenuated lipopolysaccharide-dependent mitogen-activated protein kinase p38 phosphorylation.

Conclusions PFD has lung allograft protective properties, and in addition to its known effects on T-cell biology, PFD immune-modulating activities encompass inhibitory effects on DC activation and function.

1 Division of Respiratory Medicine, Department of Medicine, Children’s Hospital Boston, Harvard Medical School, Boston, MA.

2 Department of Pathology, Children’s Hospital of Philadelphia, University of Pennsylvania, Philadelphia, PA.

This work was primarily supported by a National Institutes of Health and National Institute of Allergy and Infectious Diseases Grant R03 AI074646 and also supported by a National Heart, Lung, and Blood Institute Grant RO1 HL088191 to Gary Visner

The authors declare no conflicts of interest.

3 Address correspondence to: Gary A. Visner, D.O., Division of Respiratory Medicine, Department of Medicine, Children’s Hospital Boston, Enders 416, 300 Longwood Ave., Boston, MA 02115.


P.B. participated in making the research design, performing the research, analyzing data, and writing the article. K.L. participated in making the research design and performing the research. L.W. participated in performing the research and analyzing data. W.W.H. participated in making the research design and writing the article. G.A.V. participated in making the research design, analyzing data, and writing the article.

Supplemental digital content (SDC) is available for this article. Direct URL citations appear in the printed text, and links to the digital files are provided in the HTML text of this article on the journal’s Web site (

Received 19 January 2012. Revision requested 10 February 2012.

Accepted 30 March 2012.

Pirfenidone (5-methyl-1-phenyl-2-(1H)-pyridone) (PFD) is an orally active antifibrotic agent shown to be efficacious in fibroproliferative disease models, including pulmonary fibrosis (1–3). In clinical trials, PFD was found to have a short half-life (~4 hr) but showed limited adverse effects and was well tolerated by most patients (4, 5). Because of its safety profile and potential to ameliorate both proinflammatory and profibrotic processes, we evaluated its role in transplantation.

Our laboratory previously showed that PFD reduces obstructive airway disease in mouse heterotopic tracheal transplants and transplant-mediated fibrosis in rat lung transplants (6–8). Moreover, PFD delayed acute rejection and prolonged mouse heterotopic cardiac allograft survival (9). PFD protective properties are believed to be from its inhibition of profibrotic and inflammatory cytokines (5, 10, 11). However, we also observed a modest but significant effect on T-cell activation (9), demonstrating immune-modulating properties for PFD. Although PFD inhibition on T-cell proliferation and activation was significant, the response we observed in vivo seemed more robust and may not be explained solely by its inhibitory effect on T cells.

Dendritic cells (DCs) are professional antigen-presenting cells capable of inducing activation of both innate and adaptive immune systems. DCs sense their environment through their pattern recognition receptors and respond by secreting cytokines and chemokines, thereby activating both innate immunity and adaptive immunity. Upon exposure to foreign antigens, DCs uptake, process, and present antigens to T cells (12). Stimulation of DC receptors initiates maturation, thereby loading antigen into major histocompatibility complex(MHC) molecules, up-regulation of costimulatory molecules and MHC, and activation of T cells (13).

In this study, we further explored PFD actions on inflammation to better understand its immune-modulating properties. We hypothesized that PFD has inhibitory effects on DC activation and abrogates its capacity to induce innate and adaptive immune response. Therefore, we examined the effects of PFD and long-acting deuterated PFD (dPFD) in vivo using the mouse lung transplant model and in vitro using bone marrow–derived DCs. We observed a remarkable inhibitory effect of PFD on DC activation, maturation, and function.

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PFD Reduces Mouse Lung Allograft Rejection/Injury and DC Activation

Murine lung transplants with a full MHC mismatch, BALB/c to C57BL/6, were used to examine PFD in vivo effects on DCs. Lung allografts were initially evaluated by in vivo micro–computed tomography (μCT) imaging with mouse thorax μCTs illustrating the consolidation of untreated lung allografts, whereas PFD-treated allografts show little disease (Fig. 1A). Lung function based on peak airway pressure with delivery of a fixed tidal volume was also improved in PFD- and dPFD-treated allografts (Fig. 1B). PFD and dPFD treatment groups showed less acute cellular rejection (Fig. 1C,D), approximately A3 based on the International Society of Heart and Lung Transplantation grading scale (14) in untreated allografts as compared with a mean of 1.5 and 1.1 with PFD and dPFD, respectively (Fig. 1E).



To better assess PFD effects on DCs, we evaluated single-cell suspensions of treated and untreated lung allografts for CD45, CD11c, MHC class II, and CD86 expression based on flow cytometry 7 days after transplantation (Fig. 2A). As expected, PFD resulted in a decrease in the commonly used marker for leukocytes (CD45+) and, more importantly, reduced the percentage of DCs (CD11c+) within the CD45 population. Within CD11c+ cells, there was a reduction in MHC class II and CD86, suggesting an inhibition of DC activation/maturation. Lung DCs were better characterized by isolating CD11c+ cells from PFD and vehicle-treated lung allografts and were analyzed based on CD45+F4/80− CD11c+ MHCII+ cells gating and CD103 and CD11b surface expression. PFD treatment reduced CD103+ cells, which are believed to activate naive CD8+ T cells in the lungs (15, 16), although no difference in the percentage of CD11b+ cells (Fig. 2B) was noted. PFD reduced both CD4 and CD8 T cells, with a further reduction in the percentage of CD3+CD8+ T cells but little difference in the percentage of CD3+CD4+ T cells (Fig. 2C). Further evidence of PFD effects on the inflammatory response was a decrease in a few cytokines based on Luminex assay (see Table S1, SDC, including colony-stimulating factor (CSF)–3, interferon γ, interleukin (IL)-2, IL-1β, tumor necrosis factor (TNF)–α, regulated on activation normal T-cell expressed and secreted, IL-4, IL-5, IL-6, IL-13, inducible protein 10, and monocyte chemotactic protein (MCP)–1.



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PFD Abrogates DC Activation/Maturation

To further examine PFD effect on DC activation and maturation, in vitro studies were performed using a known activator of DCs, lipopolysaccharide (LPS) (17). As expected, LPS stimulated DC activation and maturation, whereas PFD decreased the maturation markers, MHC class II, CD80, and CD86 expression (Fig. 3A) after LPS stimulation. To determine if PFD altered DCs capacity for endocytosis, we examined the ability of these cells to uptake alloantigens. Lysates of carboxyfluorescein diacetate succinimidyl ester (CFSE)–labeled BALB/c spleen cells were incubated overnight with DCs from C57BL/6 mice, and endocytosis was measured based on CFSE uptake within DCs. No significant difference was seen in DCs uptake of allogeneic cell lysates, with or without PFD exposure, indicating that PFD does not affect DC endocytosis (Fig. 3B).



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PFD Inhibits DC–T-Cell Interaction

In transplanted tissue, donor (direct allorecognition)- and recipient (indirect allorecognition)- derived DCs stimulate T cells by presenting antigen by means of MHC class I/II and costimulatory molecules (18). To address whether PFD affects DC–T-cell interactions, we examined both direct and indirect pathways of DCs to allospecific T-cell activation based on IL-2 secretion. C57BL/6 or BALB/c bone marrow (BM)–derived DCs were initially incubated with PFD, placed in media without PFD, and exposed to previously sensitized C57BL/6–derived CD3+ T cells to stimulate an allogeneic response. As shown in Figure 4A, PFD exposure resulted in impaired T-cell stimulation with limited IL-2 secretion indicating an inhibition of DCs ability to activate T cells through direct and indirect stimulation.



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PFD Inhibits DC Cytokine Production

As another measure of DC activation, we analyzed the effect of PFD on cytokine production of LPS-stimulated DCs. PFD inhibited the secretion of CSF-3, IL-10, MCP-1, CCL12 (MCP-5), soluble TNF receptor 1, and TNF-α, whereas there was no significant change in the levels of IL-6, IL-9, IL-12p40, IL-12p70, or regulated on activation normal T-cell expressed and secreted (Fig. 4B). This data showed that PFD selectively inhibits cytokine release from stimulated DCs.

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PFD Selectively Inhibits p38 Map Kinase Phosphorylation

The mitogen-activated protein kinase (MAPK) signaling pathway is activated under a variety of cellular stresses including LPS-induced maturation and up-regulation of DC surface antigens by means of two prototype families of MAPK, p38 and c-Jun NH2-terminal kinase (JNK) (19, 20). We examined whether PFD actions may be mediated by altering p38 and JNK. In DCs, PFD attenuated LPS-stimulated p38 phosphorylation after 30 and 60 min, whereas it showed little effect on JNK-stimulated phosphorylation (Fig. 5). The relative increase of p38 phosphorylation in PFD-treated cells to LPS was significantly less as compared with non-PFD exposed cells to LPS.



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PFD is best known as an antifibrotic agent and shown to be efficacious in blocking the development of fibroproliferative disorders (1, 21). PFD was initially believed to have little significant immunosuppressive properties. However, our laboratory and others have shown that it inhibits both inflammatory and profibrotic cytokine expressions and has beneficial effects against immune-mediated injury models such as multiple sclerosis, asthma, and transplantation (6, 7, 22–24). Its immunosuppressive effects are partially related to its inhibition of proinflammatory cytokines; however, it is now clear that PFD has additional immune-modulating activities with impairment of T-cell activation, and this study demonstrated its inhibitory actions on DCs. Using an in vivo model, mouse lung transplantation showed that PFD not only reduced lung injury/rejection but also had an inhibitory effect on lung DC activation. This was also seen systemically with reduced CD11c+ cells and activation markers in the thoracic lymph nodes and spleen (data not shown).

Lung allograft DC profile was better characterized based on the two major myeloid CD11c+MHC class II+ DC populations in the lung, CD11b+CD103 and CD11bCD103+ (25). Although we observed no difference in CD11b+CD103 DCs, there was an inhibition of CD103+ DCs. Although the roles of CD103+ and CD103 DCs are not rigorous, CD103+ DCs are believed to preferentially cross-present antigen to CD8+ T cells, whereas CD103 DCs present antigen to CD4+ T cells (15, 16). This may explain why we observed a reduction in CD8+ as compared with CD4+ T cells. Interestingly, mice with a targeted disruption of CD103 were also shown to have less rejection of islet allografts (26).

In this study, PFD was given 1 day before transplantation. We found that PFD given after transplantation demonstrated limited success in suppressing transplant-mediated injury of mouse lung allografts (data not shown). We also observed this in mouse heterotopic tracheal transplants (8). That PFD needs to be given early in the transplant process suggests its primary actions may be on DC activation, thereby preventing the early innate and adaptive process rather than having a potent immune effect directly on T cells with little if any benefit as a rescue agent.

PFD has a short half-life, making delivery to mice or patients difficult (27). Recently, dPFD was developed (Patent Cooperation Treaty US2008/010565), showing a longer half-life and increased levels for the same dose. The use of deuterium does not change an agent’s biochemical potency or selectivity to pharmacological targets; however, deuterium may alter its pharmacokinetics (28). We found that dPFD at approximately one third the dose was equally if not more efficacious than standard PFD.

We used different methods to assess mouse lung allograft injury including μCT imaging, lung function, and histology. This is the first report for mouse lung allograft imaging that we are aware of. The imaging studies coincided with histology showing minimal rejection in PFD-treated allografts, whereas untreated grafts showed severe rejection. A major advantage of this technique is that mouse lung allografts can be monitored over time because it is not a terminal outcome measure as are the other techniques.

We found that PFD has an inhibitory action on stimulated DC activation, maturation, and function. This was not attributed to cell death caused by drug toxicity because, in the preliminary phase of the study, we observed a similar ratio of cell viability with or without PFD based on forward and side scatter light properties and propidium iodide (data not shown). We also found a reduced response to the Toll-like receptor (TLR)–2 and TLR 3 agonists (zymosan and polyinosinic:polycytidylic acid, respectively) after PFD treatment (data not shown). To go along with the more immature DC phenotype with PFD, DC endocytosis was not impaired by PFD. However, PFD demonstrated a remarkable inhibitory effect on DC’s ability to present antigen and stimulate T-cell activation. This was addressed for both direct and indirect pathways. As expected, we found that the direct response was more potent (18), and PFD inhibited both pathways with a nearly 50% reduction of IL-2 release for the direct pathway and a nearly complete inhibition of the indirect pathway.

PFD inhibitory effects may be attributed not only to lower MHC and costimulation expression in PFD-treated DCs but also to lower inflammatory cytokine secretion of DCs. This was evident by selective inhibition of proinflammatory cytokines from stimulated DCs in response to PFD including TNF-α, soluble TNF receptor 1, CSF-3, MCP-1 and MCP-5, and IL-10. Interestingly, we observed a decrease in the anti-inflammatory cytokine IL-10 that participates in recovery phase of infection and reduces tissue damage (29, 30). However, there was no significant decrease in IL-10 from mouse lung allografts treated with PFD (see Table S1, SDC,, and we did not see an effect on IL-10 secretion from T cells in response to PFD (9), so the overall effect of PFD on IL-10 seems to be limited.

The inflammatory response along with DC activation and cytokine production is mediated through a few signaling molecules. We initially performed a TLR signaling pathway polymerase chain reaction microarray and found that PFD inhibited several kinases including MAPK signaling molecules. Therefore, we evaluated whether PFD alters a major MAPK signaling cascade. Previously, PFD was shown to inhibit p38 and JNK gene expression in cardiac tissue (31). We also observed a 25% to 40% decrease in p38, JNK, and extracellular regulated kinase protein levels with PFD exposure of DCs (data not shown). Similar to previous studies, LPS stimulation of DCs increased p38 MAPK and JNK activation/phosphorylation (32, 33), whereas PFD inhibited p38 phosphorylation but not JNK phosphorylation. The selective inhibition of the MAPK signaling pathways could be one explanation for the differential cytokine response in response to PFD.

This study illustrates that PFD has an inhibitory affect on stimulated DCs and that this is a potential mechanism for its protection of lung allografts. Future studies are planned to more definitively substantiate the actions of PFD on DCs in transplantation by ex vivo treatment of DCs with PFD and subsequent adoptive transfer. However, this work further supports the benefits of PFD in transplantation and its potential as a therapeutic agent. Compared with most therapies for transplantation, PFD has relatively minimal adverse effects. Despite the encouraging results of this study, we do not expect PFD to be effective as a single therapy in the long term, rather we expect it to be effective as an adjunct therapy that will augment immunosuppression without a significant increase in drug toxicity. The other potential therapeutic benefit is its antifibroproliferative actions, thereby reducing fibrosis, which is a hallmark of chronic rejection.

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Lung Transplantation

BALB/c (H-2d) and C57BL/6 (H-2b) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). The Institutional Animal Care and Use Committees of the Children’s Hospital Boston approved all protocols. Orthotopic mouse lung transplantation was performed with modifications as previously described for rat lung transplantation (6, 7). Briefly, the mouse donor (BALB/c) was anesthetized with ketamine/xylazine, intubated orotracheally, and ventilated with 1% to 2% isoflurane/air at 120 breaths per min with a tidal volume of 8 mL/kg (Harvard Rodent Ventilator Model 687; Harvard Apparatus, Boston, MA). A thoracotomy was performed, and the lung was flushed with ice-cold low-potassium dextran preservation solution (Perfadex; Vitrolife, Göteborg, Sweden). The left lung was isolated, and cuffs were placed into the left pulmonary vein, bronchus, and artery. The recipient (C57BL/6) was anesthetized with ketamine/xylazine, intubated, and ventilated with isoflurane/oxygen. A thoracotomy was performed, and the donor lung was implanted using the three-cuffed hilar structures. After implantation, the incision was closed, and the recipient mouse was extubated and monitored after surgery. Seven days after transplantation, in vivo assessment of transplanted lung function was performed by measuring peak airway pressure as previously described (7). Acute cellular rejection was assessed by hematoxylin-eosin staining of lung tissue and based on grading criteria (A0–A4) from the International Society of Heart and Lung Transplantation (14).

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Experimental Groups

The experimental groups included untransplanted control (normal), isografts, vehicle-treated allografts, PFD 400 mg/kg (CAS 53179-13-8; Department of Chemistry, University of Florida, Gainesville, FL), and dPFD 150 mg/kg (Organix Inc., Woburn, MA). PFD and dPFD solutions were prepared in 10% dimethyl sulfoxide (Sigma-Aldrich, St. Louis, MO) and given by subcutaneous injection 1 day before lung transplantation and continued for 7 days after surgery.

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Computed Tomography

μCT imaging was performed using the Siemens MicroCAT II (Washington, D.C.) scanner with the radiation detector configured for a 5.4-cm transaxial field of view and an 8-cm axial field of view. Animals were anesthetized (isoflurane or ketamine/zylazine) and positioned headfirst prone on the imaging table. Data were acquired with x-ray tube voltage of 80 kilovolt (peak), tube currents of 500 μA, and exposures per view of 300 milliseconds for 200 views.

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Lung DCs

Single-cell suspensions were obtained from the lungs 7 days after transplantation by enzyme digestion (Liberase Blendzyme 3; Roche Applied Science, Indianapolis, IN). Antibodies were purchased from BD Biosciences (San Jose, CA) and eBiosciences (San Diego, CA). DC phenotyping was based on flow cytometry (LSR II; BD Biosciences) using the following antibodies: anti-CD11c, anti-CD86, anti-CD45, and anti–I-A/I-E (MHC), whereas T cells were stained for anti-CD45, anti-CD3, anti-CD4, and anti-CD8. In separate experiments, highly purified DCs were obtained by magnetic sorting of CD11c+ cells (STEMCELL Technologies, Vancouver, Canada) and stained with anti-CD103, anti-CD11b, anti-F4/80, anti-CD86, anti–I-A/I-E, and anti-CD45. Viable cells were selected based on forward and side scatter light properties and analyzed with FlowJo software (Tree Star software Inc., Portland, OR).

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DC Isolation and Maturation

C57BL/6 BM–derived DCs were generated as previously described (34). Cells were plated in Roswell Park Memorial Institute medium with 10% fetal bovine serum supplemented with penicillin, streptomycin, glutamine, and recombinant mouse granulocyte-macrophage CSF-2 (eBioscience). DCs (>95%) were obtained at day 7 by CD11c+ magnetic cell sorting (STEMCELL Technologies) and cultured with or without 2 mM PFD for 72 hr and stimulated with 1 μg/mL LPS (Escherichia coli 0127:B8; Sigma-Aldrich) for 24 hr.

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DC In Vitro Endocytosis

BALB/c allogeneic splenocytes were labeled with CFSE (Sigma-Aldrich), and cell lysates were obtained through three rapid freeze/thaw cycles. Endocytosis was measured by incubating CFSE cell lysates with DCs with or without PFD for 24 hr and washing them with phosphate-buffered saline, and the uptake of labeled cell lysates was then analyzed by flow cytometry.

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DC Antigen Presentation

DC antigen presentation in response to PFD was evaluated with modifications as previously described (35). BALB/c spleen cells were injected intraperitoneally into C57BL/6 mice, and 2 weeks later, spleen CD3+ T cells were isolated by flow cytometry sorter MoFlo (Beckman Coulter Inc., Miami, FL). The indirect pathway was analyzed by incubating C57BL/6-derived DCs with sensitized syngeneic isolated CD3+ T cells at 1:1 ratio and BALB/c allogeneic splenocyte lysates at 1:5 ratio for 24 hr. The direct pathway was analyzed by incubating T cells with BALB/c BM–derived DCs. T-cell activation was assessed by enzyme-linked immunosorbent assay for secreted IL-2 (RayBiotech Inc., Norcross, GA). Before co-culturing DCs with T cells, media were exchanged with fetal bovine serum and PFD-free media.

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Cytokine Production

C57BL/6 BM–derived DCs were cultured with or without PFD for 72 hr and stimulated with LPS, and the supernatant was analyzed after 24 hr for cytokine release by Mouse Cytokine Antibody Array (RayBiotech Inc.). The relative amounts were determined by densitometry with membrane background measurements subtracted from blanks and normalized to the positive control readings.

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p38 and JNK Phosphorylation Assay

C57BL/6 DCs with or without PFD were stimulated with LPS for 0, 10, 30, and 60 min. Intracellular p38 MAPK and JNK activation were measured using Phospho-p38 and Phospho JNK enzyme-linked immunosorbent assay (RayBiotech). To calculate p38 and JNK phosphorylation, the intensity of color was measured at 450 nm, and optical density value for each time point was normalized to 0 min.

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Data were expressed as mean±SEM, and statistical analyses were performed using the GraphPad Prism statistical program (GraphPad Software Inc., San Diego, CA). One-way analysis of variance and the Tukey multiple comparison test were used to evaluate differences between groups; and Student t test, to compare two groups. P values less than 0.05 were considered significant.

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Pirfenidone; Lung transplantation; Dendritic cell; Acute rejection; Mouse

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