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Asthma is characterized by chronic lung inflammation and airway hyperresponsiveness. Despite recent advances in understanding of its pathophysiology, asthma remains a major public health problem, and new therapeutic strategies are urgently needed. In this context, we sought to ascertain whether treatment with the TK inhibitor dasatinib might repair inflammatory and remodelling processes, thus improving lung function, in a murine model of asthma.
Animals were sensitized and subsequently challenged, with ovalbumin (OVA) or saline. Twenty‐four hours after the last challenge, animals were treated with dasatinib, dexamethasone, or saline, every 12 h for 7 consecutive days. Twenty‐four hours after the last treatment, the animals were killed, and data were collected. Lung structure and remodelling were evaluated by morphometric analysis, immunohistochemistry, and transmission electron microscopy of lung sections. Inflammation was assessed by cytometric analysis and ELISA, and lung function was evaluated by invasive whole‐body plethysmography.
In OVA mice, dasatinib, and dexamethasone led to significant reductions in airway hyperresponsiveness. Dasatinib was also able to attenuate alveolar collapse, contraction index, and collagen fibre deposition, as well as increasing elastic fibre content, in OVA mice. Concerning the inflammatory process, dasatinib reduced inflammatory cell influx to the airway and lung‐draining mediastinal lymph nodes, without inducing the thymic atrophy promoted by dexamethasone.
In this model of allergic asthma, dasatinib effectively blunted the inflammatory and remodelling processes in asthmatic lungs, enhancing airway repair and thus improving lung mechanics.
These Tables list key protein targets and ligands in this article which are hyperlinked to corresponding entries in http://www.guidetopharmacology.org, the common portal for data from the IUPHAR/BPS Guide to PHARMACOLOGY (Pawson et al., 2014) and are permanently archived in the Concise Guide to PHARMACOLOGY 2015/16 (Alexander et al., 2015).
Allergic asthma is a major global health problem and approximately 300 million people worldwide are believed to be affected by asthma. Particular negative consequences of this disease include poor health‐related quality of life, impaired work productivity, and limitations of the activities of daily living (Peters et al., 2006). Classically, CD4+ Th2 cells have been considered as the primary regulators of the allergic response through production of Th2 cytokines, which lead to airway inflammation and hyperresponsiveness (Holloway et al., 2010). Furthermore, patients with asthma may also display airway hyperresponsiveness with no early‐phase or late‐phase inflammation present, which is likely to be due to long‐term changes in the structure of the airway, such as hypertrophy and hyperplasia of the airway smooth muscle layer (Murdoch and Lloyd, 2010). Continued interaction through experimentation or modelling will be required to refine the phenotypes related to outcomes and delineate specific treatments for specific phenotypes (Chung and Adcock, 2013).
The most widely employed therapies for persistent asthma, including inhaled corticosteroids and bronchodilators (short‐lasting or long‐lasting β‐adrenoceptor agonists or muscarinic receptor antagonists), reduce the symptoms of the disease (Kim et al., 2011; Raissy et al., 2013). However, corticosteroids may not bring instant relief, thus decreasing treatment compliance by patients (Cooper et al., 2015), which accelerates disease progression and lung remodelling, leading to progressive loss of lung function (Pascual and Peters, 2005). Moreover, long‐term use of corticosteroids cannot reverse lung remodelling and may result in long‐lasting side effects, including immunosuppression, predisposing patients to lung infection with opportunistic micro‐organisms (Barnes, 2012). Finally, not all asthmatic patients benefit from inhaled corticosteroids; patients who have a more neutrophilic lung inflammatory status are particularly resistant to standard glucocorticoid anti‐inflammatory therapy (Barnes, 2013). Thus, new therapeutic approaches are needed to both attenuate inflammatory and remodelling processes in different asthma phenotypes and reduce airway resistance.
As several intracellular signalling pathways involved in inflammation and fibrosis are triggered by tyrosine kinases (Wollin et al., 2014), an intervention on this mechanism that could inhibit the inflammatory process and asthma‐associated structural changes would improve lung function. Within this context, dasatinib, a protein kinase inhibitor with main activity against BCR/ABL tyrosine kinases and broad activity against Src‐family tyrosine kinases, appears promising. This orally active, second‐generation tyrosine kinase inhibitor is primarily used for the treatment of leukaemia (Blake et al., 2008). Dasatinib also inhibits the function of some inflammatory cell types, including T lymphocytes (Schade et al., 2008), and targets a major pro‐fibrotic pathway triggered by TGF‐β (Distler and Distler, 2010). Dasatinib is also associated with the inhibition of Syk phosphorylation. Syk inhibitors are known to block the release of mediators, such as histamine, the production of PGs and leukotrienes, and the secretion of cytokines by mast cells (Gilfillan and Rivera, 2009). These properties indicate that Syk could be a novel pharmaceutical target for the treatment of allergic conditions, such as asthma.
In this study, we compared the tyrosine kinase inhibitor dasatinib with dexamethasone for its effects on inflammation and lung remodelling processes, thus improving lung mechanics, in a murine model of ovalbumin (OVA)‐challenged allergic asthma.
The protocol of study was approved by the Animal Care Committee of the Health Sciences Centre, Federal University of Rio de Janeiro, Brazil (CEUA‐019). All animals received humane care in compliance with the Principles of Laboratory Animal Care formulated by the National Society for Medical Research and the Guide for the Care and Use of Laboratory Animals prepared by the National Academy of Sciences, USA. Animal experiments have followed the ARRIVE guidelines (Kilkenny et al., 2010; McGrath and Lilley, 2015).
To produce a chronic allergic asthma model, female BALB/c mice (20–25 g) were randomly assigned to undergo sensitization and challenge with sterile OVA (Sigma‐Aldrich, St. Louis, MO, USA) or saline. In the allergic asthma groups, mice were sensitized by i.p. injection of OVA (10 μg in 0.1 mL saline) every other day for up to seven injections. Forty days after the start of sensitization, intratracheal OVA challenges were administered (20 μg in 20 μL saline), three times with 3‐day intervals between challenges (Xisto et al., 2005). The control group (CTRL group) received saline instead of OVA during both sensitization and challenge. Twenty‐four hours after the last challenge, treatment with dasatinib 1 mgkg−1 (OVA + DAS1 group) or 10 mgkg−1 (OVA + DAS10 group), dexamethasone 0.5 mgkg−1 (OVA + DEXA group), or saline (OVA group) was started and administered every 12 h for 7 consecutive days. Dasatinib and saline were administered by gavage and dexamethasone by i.p. injection.
Lung mechanics, all histological data, and mediators in lung tissue homogenate (n = 7 for each experimental group) were measured 24 h after the last treatment, as described in the following sections. Following the same protocol, a different set of animals was used for collection of bronchoalveolar lavage fluid (BALF), blood, mediastinal lymph nodes, thymus, and bone marrow data (n = 6 for each experimental group).
Twenty‐four hours after the last treatment, animals were sedated (diazepam 1 mg i.p.), anaesthetized (thiopental sodium 20 mgkg−1 i.p.), tracheotomized, and paralysed (vecuronium bromide 0.005 mgkg−1 i.v.). Lung mechanics was assessed as described previously (Olsen et al., 2011). Briefly, airway responsiveness was assessed as a change in airway function 24 h after the last treatment following aerosolized methacholine in a FinePoint R/C Buxco Platform (Buxco Electronics, Sharon, CT, USA). Airflow and transpulmonary pressure were recorded using a Buxco Pulmonary Mechanics Processing System (Buxco Electronics, Wilmington, NC, USA). Airway resistance (cm H2O.s per mL) and lung elastance (cm H2O per mL) in each breath cycle were calculated with a Buxco system as well. Analogue signals from the computer were digitized using a Buxco Analogue/Digital Converter (Buxco Electronics). Mice were allowed to stabilize for 5 min and increasing concentrations of methacholine (3, 9, 27, and 81 mgmL−1) were administered by aerosol for 5 min each. Expressed results comprised the mean absolute values of the responses collected during 5 min after administration of methacholine.
All histological analyses were performed in a blinded manner, i.e., the observer was unaware of the experimental protocol. After anaesthesia, heparin (1000 IU) was injected i.v., and the animals exsanguinated. The lungs were removed, fixed, and embedded in paraffin. Sections (4μm) were cut and stained with haematoxylin and eosin. Morphometric analysis was performed using an integrating eyepiece with a coherent system consisting of a grid with 100 points and 50 lines of known length coupled to a conventional light microscope (Axioplan, Zeiss, Oberkochen, Germany). The volume fractions of the lung occupied by collapsed alveoli (alveoli with rough or plicate walls) or normal pulmonary areas were determined by the point‐counting technique, at a magnification of ×200 across 10 random, non‐coincident microscopic fields. Briefly, points falling on collapsed or normal pulmonary areas were counted and divided by the total number of points in each field of view (Weibel, 1990).
The contraction index of the central and peripheral airways was determined by counting the points falling on the airway lumen and those falling on airway smooth muscle and on the epithelium, at a magnification of ×400. The perimeter of the airways was estimated by counting the intercepts of the lines of the integrating eyepiece with the epithelial basal membrane. The areas of smooth muscle and airway epithelium were corrected in terms of airway perimeter by dividing their values by the number of intercepts of the line system with the epithelial basal membrane of the corresponding airway. Because the number of intercepts (NI) of the lines with the epithelial basal membrane is proportional to the airway perimeter and the number of points (NP) falling on airway lumen is proportional to airway area, the magnitude of bronchoconstriction (contraction index, CI) was computed by the relationship CI = NI√NP. Measurements were performed in five airways from each animal at ×400 magnification. Collagen and elastic fibre content were quantified using a digital analysis system and image processing software (Image‐Pro Plus 5.1 for Windows; Media Cybernetics, Silver Spring, MD).
Strips (2 × 2 × 10 mm) from the right lung were fixed and embedded in paraffin for immunohistochemical analysis using a monoclonal antibody against α‐smooth muscle actin (Dako, Carpenteria, CA, USA) at a 1:500 dilution. Analysis was performed by the point‐counting technique using a 121‐point grid.
To obtain a stratified random sample, three 2 × 2 × 2‐mm slices were cut from different segments of the left lung and fixed. Ultrathin sections from selected areas were examined and micrographed in a JEOL electron microscope (JSM‐6100F; Tokyo, Japan). Submicroscopic analysis of lung tissue was performed, and pathological findings graded on a five‐point, semi‐quantitative, severity‐based scoring system.
BALB/c mice were killed with a lethal dose of pentobarbital sodium, and their tracheas were immediately cannulated. Cell influx into the airway lumen was quantified by counting cells recovered from bronchoalveolar lavage. BALF was obtained by flushing the airways three times with 0.4 mL of 1% PBS solution and retrieving the fluid by gentle aspiration. BALF was centrifuged (239 x g, 10 min) and the cell pellet re‐suspended in PBS. Cells from the lung‐draining mediastinal lymph nodes and thymus were obtained through mechanical maceration of the organs. Bone marrow cells were aspirated from the femur by flushing the bone marrow cavity with 1 mL of PBS. Total leukocyte counts were performed with Türk solution in a Neubauer chamber. Thymus weight was also measured.
Eosinophils, neutrophils, and monocytes recovered from BALF and blood were stained using a commercial kit (Panótico Rápido LB®, Pinhais, RS, Brazil). T cells recovered from BALF, mediastinal lymph nodes, and thymus were distinguished after staining with the monoclonal antibodies anti‐mouse CD3 FITC, anti‐mouse CD4 PE, and anti‐mouse CD8 PECy5 (BD). Bone marrow eosinophils and neutrophils were differentially stained with monoclonal antibodies, anti‐mouse Gr‐1 APC (eBiosciences, San Diego, CA, USA) and SiglecF PE (BD, Franklin Lakes, NJ, USA). These cells were acquired with a FACSCalibur flow cytometer (BD Biosciences PharMingen) and analysed with the FlowJo 7.6.5 software suite.
IL‐4, IL‐13, IL‐10, VEGF, eotaxin, TNF‐α (Peprotech, New Jersey, NJ, USA), and TGF‐β (Biolegend, San Diego, CA, USA) were quantified in lung tissues from all groups by elisa, performed according to the manufacturer's protocol.
Fibroblasts were isolated from lungs of control and OVA‐exposed mice. Two mice in each condition were killed by an overdose of pentobarbital. Lungs were carefully removed, cut into 1‐mm pieces, and subjected to enzymic digestion by collagenase‐1 (1 mgmL−1) for 40–60 min at 37°C. The remaining solution was pipetted up and down to break any clumps every 10 min. When media became cloudy and lung fragments changed colour from red to white and started forming sticky fibres, lung digestion was complete. We then blocked the enzymatic reaction by adding media supplemented with 10% FBS and immediately centrifuging at 524 x g for 5 min. We re‐suspended the pellet in 10 mL of warm DMEM–F12, supplemented with 10% FBS, 1000 UmL−1 penicillin/streptomycin, and 2 mM L‐glutamine (Invitrogen Life Technologies, Grand Isle, NY, USA) and then allowed the cells to settle for 24 h in a tissue culture dish. We then discarded the supernatant and changed the media every 2 days.
First‐passage cells were plated in 6‐well plates (106 cells per well) for 48 h. The medium was then replaced with fresh medium, and cells were exposed to dasatinib (100 ngmL−1 medium) or normal medium for 24 h. The supernatant was removed, cells washed with PBS, lifted using 2.5% trypsin/EDTA (Invitrogen Life Technologies, Grand Isle, NY, USA), and pelleted by centrifugation (600 x g for 5 min).
A quantitative real‐time reverse transcription PCR was performed to measure mRNA expression of Procollagen (PC) I, PC III, and TGF‐β. Cells were lysed for RNA extraction through the RNeasy Plus Mini Kit (Qiagen, Valencia, CA, USA) as per manufacturer's recommendations. The total RNA concentration was measured by spectrophotometry in a Nanodrop ND‐1000 system. First‐strand cDNA was synthesized from total RNA using an M‐MLV reverse transcriptase kit (Invitrogen). Relative mRNA levels were measured with an SYBR Green detection system using ABI 7500 real‐time PCR (Applied Biosystems, Foster City, CA, USA). All samples were measured in triplicate. The relative level of each gene was calculated as the ratio of the study gene to the control gene (acidic ribosomal phosphoprotein P0 [36B4]) and given as the fold change relative to control fibroblasts incubated with regular medium. The following PCR primers were used: Procollagen I forward: 5′‐AGA AGT CTC AAG ATG GTG GCC G‐3′ and reverse 5′‐GGT CAC GAA CCA CGT TAG CAT C‐3′; Procollagen III forward 5′‐CAG CTA TGG CCC TCC TGA TCT T‐3′ and reverse: 5′‐GTA ATG TTC TGG GAG GCC CG‐3′, TGF‐β forward: 5′‐ATA CGC CTG AGT GGC TGT C‐3′ and reverse: 5′‐GCC CTG TAT TCC GTC TCC T‐3′; 36β4 forward 5′‐CAA CCC AGT TCT GGA GAA AC‐3′ and reverse 5′‐GTT CTG AGC TCC CAC AGTGA‐3′.
These studies comply with the recommendations on experimental design and analysis in pharmacology (Curtis et al., 2015). The sample size calculation for testing the primary hypothesis (airway responsiveness is increased in OVA group) was based on previous measurements made by us (Xisto et al., 2005; Burburan et al., 2007) and on pilot studies. Accordingly, a sample size of six animals per group (considering one animal as dropout) would provide the appropriate power (1 − β = 0.8) to identify significant (α = 0.05) differences in airway resistance between OVA animals and those treated with dasatinib, taking into account an effect size d = 1.9, a two‐sided test, and a sample size ratio = 1 (G*Power 184.108.40.206, University of Düsseldorf, Germany). Statistical analyses were performed using one‐way anova or anova on ranks followed by Tukey's test or Dunn test, respectively, whereas two‐way anova followed by Tukey's test was used to evaluate lung function. Repeated measures anova was used to compare methacholine dose ranges. Parametric and non‐parametric data were expressed as mean ± SD and median (interquartile range) respectively. All tests were performed using GraphPad Prism v6.00 (GraphPad Software, La Jolla, California, USA).
Methacholine was supplied by Sigma Chemical (St. Louis, MO, USA); pentobarbital, diazepam, thiopental, and vecuronium by Cristália (Itapira, São Paulo, Brazil); dasatinib by Bristol‐Myers Squibb (New York, NY, USA), and dexamethasone by Aché (São Paulo, Brazil).
As we did not observe significant differences on parameters associated with lung inflammation, remodelling, and function between the control groups (sensitized and challenged with saline [0.9% NaCl]) treated with saline (CTRL group), dexamethasone (CTRL + DEXA group), or dasatinib (CTRL + DAS1 or CTRL + DAS10 groups), we used the control groups as a unique control for statistical comparison with the OVA groups.
Morphometric examination of the lungs of OVA‐challenged mice treated with saline (OVA group) demonstrated a significant increase in the fractional area of alveolar collapse when compared with control (CTRL) animals, in good agreement with previous observations (Xisto et al., 2005). The alveolar collapse induced by OVA was significantly mitigated in the lungs of mice treated with dasatinib at 10 mgkg−1 (OVA + DAS10 group) (Table 1). In addition, the central airway diameter was less constricted in mice in the OVA + DEXA, OVA + DAS1, and OVA + DAS10 groups than in the OVA group (Table 1).
As expected, because of the well‐established nature of OVA‐induced inflammation (Xisto et al., 2005), the total number of leukocytes in BALF, blood, mediastinal lymph node, thymus, and bone marrow was higher in the OVA group than in CTRL (Figures 1 and and2).2). The total number of leukocytes in BALF (Figure 1A), mediastinal lymph node (Figure 2A), and thymus (Figure 2B) was significantly lower in mice treated with dasatinib at 1 mgkg−1 (OVA + DAS1 group) compared with OVA. Dasatinib at 10 mgkg−1 (OVA + DAS10 group) reduced the total number of leukocytes in blood (Figure 1B), mediastinal lymph node (Figure 2A), and bone marrow (Figure 2C). Eosinophil airway influx induced by the OVA challenge (OVA group) was significantly reduced by dasatinib either at 1 mgkg−1 (OVA + DAS1 group) or at 10 mgkg−1 (OVA + DAS10 group) (Figure 1A). Interestingly, the number of eosinophils was reduced in blood only in the group treated with dasatinib at the higher dose (OVA + DAS10 group) (Figure 1B).
The T CD4 and T CD8 populations were also quantified in BALF, mediastinal lung‐draining lymph nodes, and thymus, with an increase seen in the OVA group compared with CTRL (Figures 1A and and2A2A and B). The number of CD4 T lymphocytes and CD8 T lymphocytes in BALF was only significantly reduced in the OVA + DAS1 group (Figure 1A). Animals that received dexamethasone (OVA + DEXA group) exhibited reduced thymus weight (17.6 ± 1.4 mg ), compared with OVA animals (46.9 ± 5.5 mg) and controls (44.2 ± 2.3 mg), compatible with the immunosuppressive profile of this corticosteroid (Frawley et al., 2011). Dexamethasone (i.p.) also reduced T CD4 and T CD8 cell populations in mediastinal lung‐draining lymph nodes and thymus (Figure 2A and B). Although dasatinib inhibited inflammation in the lungs and decreased the number of T cells in the thymus and mediastinal lymph nodes (OVA + DAS1 group) induced by the OVA challenge, it did not induce as dramatic a decrease in T cells in the mediastinal lymph nodes (Figure 2A) and thymus (Figure 2B) as observed with dexamethasone treatment, compared with the CTRL group.
Furthermore, the number of neutrophils was efficiently reduced in BALF (Figure 1A) and blood (Figure 1B) by 10 mgkg−1 of dasatinib (OVA + DAS10 group). Although dasatinib did not alter the number of neutrophils in the bone marrow, dexamethasone (OVA + DEXA group) was able to significantly decrease the number of neutrophils in the bone marrow (Figure 2C).
We also measured levels of mediators crucially involved in asthma pathogenesis (Abreu et al., 2013). IL‐4 (Figure 3A), IL‐13 (Figure 3B), VEGF (Figure 3D), TGF‐β (Figure 3E), eotaxin (Figure 3F), and TNF‐α (Figure 3G) in BALF were significantly higher in the OVA group, compared with the CTRL animals. Dasatinib at 10 mgkg−1 reduced levels of these mediators and significantly increased the IL‐10 level (Figure 3C). However, only dasatinib at 1 mgkg−1 reduced TGF‐β (Figure 3E), eotaxin (Figure 3F), and TNF‐α (Figure 3G) levels. Dexamethasone (OVA + DEXA group) was efficient at reducing only IL‐4 (Figure 3A), IL‐13 (Figure 3B), and TGF‐β (Figure 3E) levels.
Hypertrophy and/or hyperplasia of airway smooth muscle associated with increased smooth muscle‐specific actin are important hallmarks of airway remodelling (Bentley et al., 2009). Interestingly, we observed a reduction in smooth muscle‐specific actin content in all treated groups (OVA + DEXA, OVA + DAS1, and OVA + DAS10), compared with the OVA group (Figure 4A).
Elastic fibre content in the alveolar septa and airways was significantly decreased in the OVA group compared with CTRL group (Figure 4B and C). We did not observe significant differences between OVA + DEXA, OVA + DAS1, and OVA (Figure 4B and C). There was a significant increase in the amount of elastic fibre in the alveolar septa and airways only in the group treated with dasatinib at the higher dose (OVA + DAS10) compared with the OVA group (Figure 4B and C).
Overproduction and deposition of extracellular matrix particularly collagen, is the most notable pathological alteration of airway remodelling (Lu et al., 2014). As expected, the collagen fibre content in the airways and alveolar septa was higher in the OVA group compared with the CTRL group (Figure 5A). Collagen fibre content was significantly reduced in the airways in OVA + DAS1 and OVA + DAS10 animals when compared with OVA (Figure 5A), but not in the alveolar septa; the latter was significantly reduced only in the OVA + DAS10 group as compared with the OVA group (P = 0.0125).
After observing the major effect of dasatinib on reducing collagen fibre content, we performed an in vitro study to analyse important fibrogenic factors in the fibroblasts. Remarkably, we observed that dasatinib (100 ngmL−1 medium) was able to reduce expression of Procollagen I (Figure 5B), Procollagen III (Figure 5C), and TGF‐β (Figure 5D) mRNA to CTRL levels.
Next, we performed transmission electron microscopy (TEM) on airway sections from animals in each group (Figure 6). Mice in the OVA group exhibited considerable subepithelial fibrosis, smooth muscle hypertrophy and hyperplasia, and mitochondrial damage (Figure 6A–C), all characteristic features of the lung remodelling present in patients with asthma (Durrani et al., 2011).
Based on a semi‐quantitative analysis of TEM images, mice in the OVA group exhibited significant basement membrane thickening, smooth muscle cell hypertrophy and hyperplasia, collagen deposition, mitochondrial damage, and endoplasmic reticulum enlargement (Table 2). Remarkably, dexamethasone (OVA + DEXA group) and dasatinib at both tested doses (OVA + DAS1 and OVA + DAS10 groups) attenuated some of these ultrastructural changes that resulted from lung remodelling. Nonetheless, treatment with dasatinib at the 10 mgkg−1 dose (OVA + DAS10 group) appeared to be more effective in this model (Table 2).
Lung elastance is expected to be elevated in asthmatic lungs undergoing a fibrotic process (Abreu et al., 2011). We found that lung elastance was higher in OVA mice than in controls (Figure 7A). This parameter was significantly attenuated in the OVA‐challenged mice treated only with dasatinib at both doses (OVA + DAS1 and OVA + DAS10 groups), but not with dexamethasone (OVA + DEXA group). Furthermore, the increase in airway resistance elicited by methacholine was significantly augmented in the OVA group (Figure 7B). However, these changes were mitigated in the groups treated with dexamethasone (OVA + DEXA group) and dasatinib at both doses (OVA + DAS1 group and OVA + DAS10 group).
Current therapies for allergic airway diseases are limited to alleviating disease symptoms and exacerbations and are unable to repair lung damage that occurs during the natural history of disease, highlighting the need for development of new therapeutic strategies (Manuyakorn et al., 2013). Dasatinib has proven value in the treatment of cancer (Kantarjian et al., 2006) and also exhibits anti‐inflammatory effects (Kneidinger et al., 2008). However, this is the first study to demonstrate that oral administration of dasatinib can mitigate airway inflammation and lung remodelling, leading to improved pulmonary mechanics in a murine model of asthma, without decreasing lymphocyte, eosinophil, and neutrophil production. We also confirmed through histological and functional findings that neither dasatinib nor dexamethasone induced any significant alterations in the lungs of normal inbred mice.
The inhibitory effects of dasatinib on T cells by the interruption of T cell receptor signalling in a dose‐dependent and time‐dependent manner have been widely investigated (Blake et al., 2008; Fei et al., 2008; Fei et al., 2009; Nerreter et al., 2013). Accordingly, we demonstrated that oral administration of dasatinib in OVA‐challenged animals inhibited the total leukocyte count in BALF, mediastinal lymph nodes, thymus, blood, and bone marrow, as well as CD4+ and CD8+ T cell influx into the bronchoalveolar space, thymus and lung‐draining lymph nodes. It is noteworthy that the inhibitory effects of dasatinib on T cells were not accounted for by toxicity (Blake et al., 2008) or apoptosis (Nerreter et al., 2013). Therefore, dasatinib inhibition of T cell accumulation in the lungs is most likely to be related to inhibition of the activation of these cells through T cell receptors and reduction of pro‐inflammatory stimulus that would lead to an increase in T cell development in the thymus.
Futosi et al. demonstrated that ex vivo adhesion of mouse peripheral blood neutrophils was strongly reduced after p.o. administration of 5 mgkg−1 of dasatinib (Futosi et al., 2012). Those results suggest that dasatinib treatment may affect the proinflammatory functions of mature neutrophils and raise the possibility that dasatinib‐related compounds may provide clinical benefit in neutrophil‐mediated inflammatory diseases. In accordance with these findings, we found that, at 10 mgkg−1 (OVA + DAS10 group), dasatinib strongly reduced neutrophil count in BALF and peripheral blood, compared with the OVA‐challenged group, without causing alterations in the bone marrow. It is important to state that dasatinib did not reduce neutrophil counts in bone marrow, lungs, or blood, compared with the CTRL group. In contrast, dexamethasone (OVA + DEXA group) significantly reduced neutrophil levels in the bone marrow, compared with the OVA and CTRL groups. Although dasatinib at the higher dose level, did reduce total leukocyte numbers in the blood and bone marrow, this effect did not appear to be related to a decrease in eosinophil or neutrophil levels. Other cells, such as monocytes, might be the target of dasatinib at the bone marrow. However, there were no significant changes in monocyte numbers in the bone marrow when dasatinib‐treated groups were compared with the non‐treated, OVA group.
Furthermore, a number of different signalling molecules have been implicated in asthmatic inflammatory changes and lung remodelling, such as IL‐13, IL‐4, eotaxin, and VEGF, levels of which increased significantly in the OVA group in this study, as previously described (Abreu et al., 2013). In our experiment, dasatinib, significantly reduced IL‐13, IL‐4, eotaxin and VEGF levels. These findings are in line with earlier investigations demonstrating that dasatinib can indeed block the production/secretion of IL‐4 as well as the VEGF signalling pathway (Kneidinger et al., 2008; Noy et al., 2012).
Another important inflammatory mediator that has been implicated in many aspects in asthma, such as recruitment of neutrophils and eosinophils and proliferation of myofibroblats, is TNF‐α (Brightling et al., 2008). Our findings also demonstrate that dasatinib is efficient at reducing TNF‐α levels.
In addition to inhibiting inflammation, dasatinib is also able to target one of the major pro‐fibrotic pathways – TGF‐β‐signalling – thus inhibiting collagen fibre synthesis (Distler and Distler, 2010). We observed a significant reduction in collagen fibre content in OVA‐challenged animals treated with dasatinib, mostly at the 10 mgkg−1 dose (OVA + DAS10 group). There was also a reduction in TGF‐β and Procollagen I and III expression in cultured fibroblasts from OVA‐challenged mice exposed to dasatinib when compared with DMSO‐exposed cells, which is consistent with previous reports (Distler and Distler, 2010). In addition, dasatinib also reduced TGF‐β levels in lung tissue homogenates.
Another important finding of this study is that dasatinib was able to mitigate the mitochondrial damage observed in OVA‐challenged mice. In this context, many researchers have been studying the role of mitochondria in the pathogenic mechanisms of asthma (Mabalirajan et al., 2008; Mabalirajan and Ghosh, 2013; Reddy, 2011). An imbalance between Th1 and Th2 immune response is crucial for the development of the pathophysiological features of asthma. A Th2‐dominant response produces oxidative stress in the airways and is thought to be one of the crucial components of mitochondrial damage and asthma pathogenesis. Although mitochondria play a crucial role in endogenous production of reactive oxygen species and mitochondrial damage and changes in their key functions are associated with experimental allergic asthma, mitochondrial involvement in this process remains unexplored (Mabalirajan et al., 2008; Wiegman et al., 2015).
This present study also compared the effects of dasatinib versus those of dexamethasone in OVA‐challenged mice. Although dexamethasone is a mainstay of anti‐inflammatory therapy for asthma, many patients with moderate to severe asthma have poor control and persistent inflammation despite high corticosteroid doses (Stirling and Chung, 2001; Bousquet et al., 2010). This suggests that inhaled corticosteroids exert sub‐optimal anti‐inflammatory effects on immune cells in the airways of patients with moderate to severe asthma. According to our results, unlike dasatinib, dexamethasone treatment induced thymic atrophy in OVA‐challenged mice. Thus, dasatinib may inhibit lung inflammation without eliciting immunosuppression, another advantage over glucocorticoid treatment.
Some limitations of the present study should be recognised. First, our results cannot be extended to other experimental models of asthma or directly extrapolated to the clinical setting. Second, only one dose of dexamethasone (0.5 mgkg−1 every 12 h) was used, based on pilot studies and on a previous study (Campos et al., 2006) that demonstrated reduced lung inflammation and improved lung function. Third, dasatinib may have side‐effects, depending on tolerability and dose level, including neutropenia, myelosuppression, and pulmonary arterial hypertension. Neither neutropenia nor myelosuppression was observed in the present study, regardless of dose. Although pulmonary arterial pressure was not measured, no vascular morphological changes suggestive of pulmonary arterial hypertension were observed.
In conclusion, in our murine model of experimental allergic asthma, dasatinib effectively blunted the inflammatory and remodelling processes in the lungs, enhancing airway repair and thus improving lung mechanics. Our results may be regarded as a very important first step, though many more studies are required to elucidate the mechanisms of action of dasatinib in asthma and enable translation to clinical research.
A.L.S., F.F.C., M.M.M., M.A.M., P.C.O., and P.R.M.R. participated in study design. A.L.S., R.F.M., V.C.B., J.D.S., F.F.C., P.S.M., and T.P.T.F. conducted the experiments. A.L.S., R.F.M., V.C.B., J.D.S., F.F.C, P.S.M. M.M.M., T.P.T.F. M.A.M., P.C.O., and P.R.M.R. performed data analysis. All authors wrote or contributed to the writing of the manuscript.
The authors declare no conflicts of interest.
This Declaration acknowledges that this paper adheres to the principles for transparent reporting and scientific rigour of preclinical research recommended by funding agencies, publishers and other organisations engaged with supporting research.
The authors would like to express their gratitude to Mr Andre Benedito da Silva for animal care, Miss Priscila Carneiro for her skilful technical assistance during the experiments, Mrs Ana Lucia Neves da Silva for her help with microscopy, Mrs Moira Elizabeth Schöttler and Mr Filippe Vasconcellos for their assistance in editing the manuscript; and Prof. Ronir Raggio Luiz, PhD (Institute of Public Health Studies, Federal University of Rio de Janeiro) for his help with statistics.
This study is funded by the Brazilian Council for Scientific and Technological Development (CNPq), the Carlos Chagas Filho Rio de Janeiro State Research Foundation (FAPERJ), the Coordination for the Improvement of Higher Level Personnel (CAPES), and the European Community's Seventh Framework Programme under grant agreement no. HEALTH‐F4‐2011‐282095 (TARKINAID project) and HEALTH‐F4‐2011‐281608 (TIMER project).
da Silva A., Magalhães R., Branco V., Silva J., Cruz F., Marques P., Ferreira T., Morales M., Martins M., Olsen P., and Rocco P. (2016) The tyrosine kinase inhibitor dasatinib reduces lung inflammation and remodelling in experimental allergic asthma. British Journal of Pharmacology, 173: 1236–1247. doi: 10.1111/bph.13430.