HB-EGF–Promoted Airway Smooth Muscle Cells and Their Progenitor Migration Contribute to Airway Smooth Muscle Remodeling in Asthmatic Mouse

Airway remodeling is characterized by structural changes in the airway walls that limit airflow in patients with chronic asthma. One of the dominant structural changes is the increase of airway smooth muscle (ASM) mass, which has been shown to be the most crucial factor that correlates with decreased lung function in asthma (1). Besides proliferation and increased size of ASM cells, the proposed mechanisms of ASM mass increase also included enhanced cell migration (2). Significant research evidence supported that ASM cells do not only migrate in the muscle bundle to induce ASM mass thickening, but also migrate through the potentially antiproliferative extracellular matrix to appear in subepithelial tissues. Furthermore, in addition to the local ASM cells migration, recent studies led us to consider the possibility of circulating CD34⁺ progenitor cells from bone marrow (also known as progenitor for smooth muscle cells) to be another source of excess ASM mass (3), because these cells were observed to be involved in the pathogenesis of smooth muscle layer thickening in vascular remodeling disease such as atherosclerosis and neointimal lesions (4) and a response of circulating progenitor cells migrating to injured lung exists in asthma cases (5, 6). Although these structural changes are known to cause substantial airflow limitation in asthma, they cannot be reversed by currently available asthma therapies. Thus, there is an urgent need to identify the novel triggers and mechanisms of ASM migration.

Several studies have shown that the expression of heparin-binding epidermal growth factor (HB-EGF), a member of the EGF superfamily (7), is increased in asthmatic airway tissues. HB-EGF can stimulate p38 and ERK1/2 MAPK (8), which are located downstream of epidermal growth factor receptor (EGFR), to play a role in ASM cells proliferation (9). Hassan et al. (10) reported the upregulation of HB-EGF expression within the asthmatic ASM mass and considered it to be a potential biomarker for the active stage of ASM remodeling. Our previous studies also found that the level of HB-EGF expression was correlated to the ASM mass thickening, and it induced ASM proliferation in vitro (11), which indicated the involvement of HB-EGF in the development of airway remodeling of asthma.

In addition to being a mitogen, HB-EGF is also a potent chemotactic factor for a number of cells including epithelial cells, fibroblasts, mesenchymal stem cells, and smooth muscle cells (7). In vascular smooth muscle (VSM) cells, it has been suggested that HB-EGF is more potent than EGF to stimulate cells migration because of its fairly strong propensity to bind cell-surface heparin-sulfate proteoglycans (12). Recently, the study by Hirota et al. (13) indicated that HB-EGF might play a role in ASM cells migration. However, this migration process and the underlying mechanisms were not fully explored. In this study, we explored the role of HB-EGF in ASM cell migration and remodeling in asthma and the underlying mechanisms.

In the current study, we established an OVA-induced asthmatic mice model and noticed that rHB-EGF could promote ASM layer thickening. Boyden chamber assay revealed that rHB-EGF facilitated ASM cells migration, with an enhanced migratory response in asthmatic ASM cells compared with their normal counterparts. p38-dependent actin cytoskeleton remodeling was involved in this rHB-EGF–induced migration process. Additionally, we detected that rHB-EGF can induce smooth muscle progenitor cells migration from circulation to asthmatic airway. Overall, our research emphasized the significant involvement of HB-EGF in ASM migration and remodeling in asthma.

Male C57BL/6 mice (4–6 wk of age) were purchased from National Rodent Laboratory Animal Resources, Shanghai Branch, and housed under specific pathogen-free conditions. The experiments were performed according to protocols approved by the Animal Studies Committee of China.

All mice were sensitized using 25 μg OVA (Grade V; Sigma-Aldrich, St. Louis, MO) in 0.1 ml aluminum hydroxide i.p. on days 0 and 12. The experimental group was challenged with aerosolized 5% OVA for 30 min daily between days 18 and 23 and euthanized on day 24 to provide an asthmatic model (11).

The rHB-EGF (2 μg/mouse; Prospec, Ness-ziona, Israel) was dissolved in PBS and intratracheally instilled in an asthmatic or normal mice every other day from days 18 to 23, 30 min before nebulization. PBS control mice received PBS only according to the same volume and schedule. Mice were sacrificed on day 24, and lung sections were stained for ASM. In another series of experiments, asthmatic or naive mice were transferred with smooth muscle progenitor cells (1 × 10⁶/mouse), which was labeled with PKH-26 red fluorescence (for flow cytometric analysis; Sigma-Aldrich) or vivotrack 680 (for in vivo optical imaging analysis; PerkinElmer, Waltham, MA) via tail vein on days 26 and 29, and intratracheally instilled with HB-EGF/PBS simultaneously. The PBS control group was instilled with PBS only. Moreover, the mice continued to receive OVA challenge three times a week after day 24 and were sacrificed on day 30.

In order to determine the thickness of ASM mass, the lung sections were incubated overnight at 4°C with the primary mAb directed against α-smooth muscle actin (α-SMA; Sigma-Aldrich). Immunoreactivity was detected by sequential incubations of lung sections with a, HRP-conjugated secondary Ab and followed by diaminobenzidine staining. The area of α-SMA immunostaining was out lined and quantified to evaluate the thickness of the ASM layer using the image analysis system (Image-Pro Plus 6.0). The results were expressed as the area of staining per micrometer length of the basement membrane of bronchioles with a 150–200-μm internal diameter at comparable sites of each slide. At least 10 bronchioles were evaluated per slide (14).

ASM cells were obtained as previously described (15). Right after that, the tracheae of asthmatic mice or normal controls were excised, washed, and digested with 0.2% collagenase IV (Sigma-Aldrich) and 0.05% elastase (Sigma-Aldrich) for 30 min at 37°C. The tissue was allowed to stand, and supernatant was collected and centrifuged (500 × g, 6 min). The pellet containing ASM cells was resuspended in 1:1 DMEM/Ham’s F12 (PAA Laboratories, Piscataway, NJ) and then plated in six-well plates. Cells at passages 2–4 were used for the following experiments.

Whole-cell lysates from ASM cells were boiled and separated by SDS-PAGE (12% SDS tricine gel) before transfer to polyvinylidene difluoride membranes. Membranes were blocked and then incubated with anti–p-EGFR (Tyr¹¹⁷³; Cell Signaling Technology, Beverly, MA), anti-EGFR (Cell Signaling Technology), anti–p-ErbB₂ (Y1248; Abcam), anti-ErbB₂ (Abcam), anti–p-p38 (Cell Signaling Technology), anti-p38 (Cell Signaling Technology), anti–p-ERK1/2 (Cell Signaling Technology), or anti-ERK1/2 (Cell Signaling Technology) overnight at 4°C. After incubation with HRP-conjugated secondary Ab (Santa Cruz Biotechnology, Dallas, TX), membranes were washed extensively, and antigenic bands were visualized by ECL (Thermal Scientific, Waltham, MA) according to the manufacturer’s protocol.

The migration experiments were performed using a 6.5-mm Transwell culture plate with an 8.0-μmol pore polycarbonate membrane (Corning). The selected cells were starved in DMEM with 0.2% BSA for 12 h. After starvation, the cells were harvested and plated on the upper side the membrane, some of which were pretreated with AG1478 (EGFR inhibitor; Calbiochem, Vienna, Austria), PD98059 (ERK inhibitor; Calbiochem), or SB203580 (p38 inhibitor; Calbiochem) separately for 30 min. The lower compartment contained the putative chemokines (rHB-EGF 10–50 ng/ml or positive control platelet-derived growth factor [PDGF]-BB). After 24 h of incubation at 37°C, the cells on the upper face of the membranes were scraped, and the cells that migrated to the lower face of the membrane were stained with crystal violet. The number of migrated cells on the lower face of the filter was counted in four random fields under ×20 original magnification (Olympus IX71; Olympus). Assays were done in duplicate using tissues from six to eight different lung specimens for accuracy of results.

Bone marrow mesenchymal stem (BMMS) cells were isolated and cultured as previously described (16). In short, bone marrow was flushed out from the femurs and tibias of mice using a 30-gauge needle and then passed through a 70-μm nylon mesh cell strainer (BD Biosciences). Cells collected after centrifugation (500 × g for 10 min) were cultured in a hypoxia chamber (Stemcell Technologies, Vancouver, Canada) using medium specialized for mouse mesenchymal stem cells (Stemcell Technologies). Cells in passage 1 were reseeded on fibronectin-coated six-well plates, and a high amount of PDGF-BB (50 ng/ml; Invitrogen, Grand Island, NY) was added to the medium for its differentiation into the smooth muscle progenitor cells (17).

Cells cultured in fibronectin-coated six-well plates were collected and resuspended for surface staining with PE-labeled anti-CD34, permeabilized with Cytofix/Cytoperm (eBioscience, San Diego, CA), and intracellularly stained with FITC-labeled anti–α-SMA. Isotype-matched IgG Abs (BioLegend, San Diego, CA) were used as negative control. Flow cytometry acquisition was performed using an FACScalibur (BD Biosciences), and results were analyzed with CellQuest software (BD Biosciences).

In another series of experiments, the mouse tracheae and lungs were prepared for single-cell suspension, and intracellular stained with FITC-labeled anti–α-SMA. Flow cytometry was used to recognize PKH-26 red fluorescence labeled cells and analyzed their concentration in α-SMA⁺ cells.

Twenty-four hours after tail vein injection, mice were anesthetized and placed in a light-sealed chamber. An in vivo optical imaging system (Caliper IVIS kinetic; Caliper Life Sciences) was used to detect Vivotrack 680–labeled cells at wavelengths 680/700 nm excitation/emission in anesthetized mice.

ASM cells were synchronized and pretreated for 12 h. After scraping by a sterile 1-ml pipette tip, the cells were washed with PBS and then treated with rHB-EGF (50 ng/ml) in the presence or absence of heparinase III (Sigma-Aldrich), AG1478, or SB203580 pretreatment. After 24 h, the cells that migrated into the scratch area were fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton X-100 for 5 min, and incubated with 5% BSA/PBS for 30 min. The cells were immunolabeled with mAbs against vinculin (Sigma-Aldrich) and followed by the Alexa Fluor 594–conjugated secondary Ab (Invitrogen). The cells were also stained with FITC-phalloidin (Sigma-Aldrich) and DAPI (Sigma-Aldrich). Images were captured with a microscope (DP12; Olympus) and a SPOT CCD camera and processed with ImagePro Plus 6.0. The polymerized actin was quantified as the percentage of polarized cells, and the focal adhesions were analyzed by the numbers of punctuate vinculin spots in 100 cells.

Data were expressed as means ± SD with a group size of six from four different experiments. Statistical analysis was performed using SPSS 16.0 (SPSS, Chicago, IL). Data between groups was compared by using one-way ANOVA followed by the Tukey honest significant difference test. A p value < 0.05 was considered statistically significant.

In our previous studies, we have demonstrated that airway epithelial expression of HB-EGF was significantly elevated following prolonged OVA challenge, and the administration of anti–HB-EGF mAb obviously attenuate OVA exposure-induced ASM mass thickening in an asthmatic murine model (11). In the current study, our results showed that airway instillation of HB-EGF to normal mice could not induce the thickening of ASM mass; however, its instillation to asthmatic mice could significantly accelerate the thickening of ASM layer compared with PBS controls (Fig. 1), which indicated the involvement of HB-EGF in the progression of ASM remodeling.

The ASM remodeling includes proliferation, hyperplasia, and migration, in which we have previously demonstrated that HB-EGF can promote ASM cells proliferation (11). In the present research, we went on to test its ability on stimulating ASM cell migration using a Boyden chamber assay. Results showed that rHB-EGF can stimulate normal ASM cell migration in a dose-dependent manner, with the maximal effect at 50 ng/ml (Fig. 2A). The ASM cells obtained from an asthmatic mouse were also tested, and a similar dose-dependent manner was observed. However, rHB-EGF was able to induce significant migration of asthmatic ASM cells at a lower concentration (20 ng/ml) than it did to normal cells (Fig. 2B), indicating the enhanced migration ability of asthmatic ASM cells.

To determine the downstream pathway of HB-EGF, the four known ErbB receptors in ASM cells were first analyzed by Western blot. Results showed that ASM cells can synthesize EGFR (ErbB₁) and ErbB₂, as shown in Fig. 3, but not ErbB₃ and ErbB₄. Furthermore, rHB-EGF can induce phosphorylation of EGFR and ErbB₂, with maximal phosphorylation occurring at 5–10 min. We also found that EGFR was significantly expressed on asthmatic ASM cells than their normal counterparts, similar to what previous studies showed (Fig. 3).

Both ERK1/2 and p38 are downstream kinases of EGFR, which are known to mediate signaling pathway related to cell migration. As expected, rHB-EGF stimulated the phosphorylation of ERK1/2 and p38 in a time-dependent manner, with maximal phosphorylation occurring at 10 min. We also used the specific EGFR inhibitor AG1478 to examine whether rHB-EGF–induced activation of ERK1/2 and p38 was EGFR dependent and found that the phosphorylation of ERK1/2 and p38 was blocked by AG1478 treatment before rHB-EGF stimulation (Fig. 3).

To ensure that the ERK1/2 and p38 pathway was actually involved in HB-EGF–induced migration, the ERK1/2 inhibitor (PD98059), the p38 inhibitor (SB203580), and AG1478 were used in the Boyden chamber system to block each pathway, respectively. Fig. 4 showed that AG1478 and SB203580 attenuated rHB-EGF–induced ASM cells migration significantly, whereas PD98059 had no effect during this process, revealing the essential roles for EGFR/p38 pathway in mediating HB-EGF–induced ASM cell migration.

Because effective reorganization of the actin cytoskeleton is absolutely important for migration, we attempt to determine whether HB-EGF can interfere with the cytoskeleton and focal adhesions. Immunofluorescent staining with FITC-phalloidin and anti-vinculin Ab of the cells migrated into the scratch area revealed that rHB-EGF induce disassembly of actin stress fibers and the formation of focal adhesions, which can be blocked by pretreatment of heparinase III, an enzyme that can cleave heparin sulfate on the ASM cell surface, and AG1478, which means that the interaction of rHB-EGF with heparin-sulfate proteoglycan or EGFR on ASM cells did play a role in the actin-mediated migration. Furthermore, the addition of SB203580 before rHB-EGF stimulation largely abolished the reorganization of actin filaments, although focal adhesion formation can still be observed, suggesting the crucial role of p38 in rHB-EGF–mediated actin stress fiber disassembly (Fig. 5).

Previous studies have demonstrated that long-term culture with PDGF-BB can induce BMMS cells differentiated into smooth muscle progenitor cells that express smooth muscle–specific markers such as α-SMA, calponin, and myosin H-chain and are also positive for CD34 and Flt1 expression (4, 17, 18). We cultured mouse BMMS cells on a fibronectin matrix with high amounts of PDGF-BB–enriched (50 ng/ml) medium. After several days of cell culture, the BMMS cells acquired a spindle-shaped phenotype and a hill and valley–like appearance. Flow cytometric analysis showed that CD34 or α-SMA was first separately expressed on a small number of cells on day 7 and for 2- to 3-wk culture, ∼70% cells were double positive for α-SMA and CD34 (Fig. 6), which could be used in the following experiments.

In order to investigate whether HB-EGF can induce smooth muscle progenitors migration in vivo, we labeled the cells with PKH-26 red fluoresces or Vivotrack 680 and employed in vivo optical imaging and flow cytometry to trace it in vivo. Twenty-four hours after transfer of cells into the tail vein, imaging techniques showed the red fluorescence emission from airways of asthmatic mice, whereas only little fluorescence was noted in naive mice. Airway instillation with HB-EGF to normal mice could hardly promote the fluorescence emission, but its administration to asthmatic mice obviously elevated the fluorescent cells’ recruitment to airways compared with the PBS group (Fig. 7A). Flow cytometry analyzed the proportion of PKH-26–labeled cells in α-SMA⁺ cells obtained from tracheae and showed almost similar results (Fig. 7B), indicating that HB-EGF instillation intratracheally might induce smooth muscle progenitor cell recruitment to the airways.

Much research evidence has supported the involvement of HB-EGF in airway remodeling of asthma, especially in the progression of ASM remodeling. HB-EGF has been demonstrated to promote ASM cell proliferation in vitro and might be a biomarker for ASM mass increase (9, 10). Our previous and present studies found that HB-EGF instillation intratracheally obviously accelerated ASM layer thickening, whereas the addition of HB-EGF Ab ameliorated OVA-induced ASM mass thickening, which proposed further evidence for the involvement of HB-EGF in ASM mass increase. However, we found that the airway instillation of HB-EGF to naive mice had no effect on the thickening of ASM mass. We considered that initiating the process of ASM migration or proliferation in asthma may need dozens of mediators, and HB-EGF might be one of them to further accelerate the ASM remodeling.

An intensive crosstalk between the epithelium and ASM has been recognized, which involves lots of mediators secreted by airway epithelium (19, 20). Although ASM cells can also secrete HB-EGF, immunohistochemical analysis has indicated airway epithelial cells to be an important and quantitatively greater biological source for the overexpression of HB-EGF in asthma (9). Given the fact that ASM cells express ErbB₁ and ErbB₂, which can form a high-affinity receptor for HB-EGF (21), HB-EGF might be an important mediator of the interaction between airway epithelial cells and ASM cells.

While investigating the effect of HB-EGF on asthmatic ASM cells versus normal ASM cells migration, we observed exaggerated migratory response in asthmatic ASM cells, although the maximal degree of migration of asthmatic ASM cells was almost similar to the normal ASM cells. This enhanced migration response to HB-EGF may be due to the increase in receptor expression, which has also been observed in our study, and sensitivity, binding capacity, and recycling in asthmatic ASM cells. In addition, the upregulated motility of the asthmatic ASM cells themselves might be another cause of interest. Wei et al. (22) observed an increase in F-actin and anti-tubulin of ASM cells obtained from asthmatic rats, which was also shown in the study by McVicker et al. (23). These changes in the actin cytoskeleton may explain their heightened migration responsiveness toward external agonists.

It is well known that EGFR signaling is followed by the activation of the MAPK pathway, including p38 and ERK, which is involved in HB-EGF–induced ASM cell proliferation (8, 13). Our present study showed similar results in that HB-EGF could stimulate the phosphorylation of p38 and ERK in ASM cells. However, for smooth muscle migration, we found that only the signal from p38 was transmitted to the downstream system to induce migration. There are conflicting views on which subfamily of MAPKs is involved in the intracellular signal pathway to induce cell migration. Similar observations have been reported for ASM cell migration induced by CXCL2 (24) or in the process of VSM cell migration (25, 26), suggesting an important role for p38 rather than ERK in cell migration. In contrast, there are also some studies that reported the involvement of ERK in ASM and VSM cell migration (27–29). Indeed, this signaling bias has been reported to occur when a ligand selectively binds and favors a specific confirmation of receptors leading to activation of specific signaling pathways (30).

Much work supports the participation of p38 MAPK in asthmatic ASM mass increase (31). Various stimuli including PDGF-BB, IL-17A/F, and CXCL2 induced ASM cell migration involved in p38 activation (24, 32, 33). In addition, the current study shows that p38 plays an important role in HB-EGF–induced actin stress fiber disassembly. In accordance with our results, studies conducted in VSM cells also showed that p38 is involved in cell migration by remodeling the actin cytoskeleton (25). Pichon et al. (34) showed that the p38/MK2/Hsp27 cascade controls actin polymerization and lamellipodium formation in PDGF-induced VSM cell migration. Esfandiarei et al. (35) observed that p38 may control actin reorganization partly via regulating the dephosphorylation and consequent activation of cofilin, a small actin-binding protein that can depolymerize and sever pre-existing actin filaments. Moreover, p38 has been demonstrated to activate the phosphorylation of caldesmon, an actin-binding protein, to induce the severing of actin filaments (36, 37).

In addition to ASM cells, we also investigated the involvement of smooth muscle progenitor cells migration in vivo. Previous studies have already found a response of BMMS cells in the lung to injury perpetrated by inflammation observed in asthma (6), and circulating CD34⁺ progenitor cells migrating to lung tissue were indeed involved in the airway remodeling (5, 38). Recently, Wu et al. (39) showed that CD34⁺ smooth muscle progenitor cells were increased in peripheral blood of asthmatic patients and involved in airway remodeling. In our study, we also discovered that OVA sensitization and challenge obviously promoted the migration of smooth muscle progenitor cells to airways and lungs compared with sham controls. Moreover, although the elevated concentration of HB-EGF was not enough to recruit smooth muscle progenitor cells, HB-EGF instillation combined with OVA treatment could significantly promote the recruitment of smooth muscle progenitors to the tracheae compared with instillation of PBS, whereas the progenitor cells migrated to the lungs were almost similar after receiving HB-EGF or PBS intratracheally as analyzed by flow cytometry (data not shown). The possible reason might be that the expression of HB-EGF in lung was significantly elevated after OVA exposure so the airway instillation of HB-EGF obviously raised the HB-EGF concentration in tracheae but not in the lungs. Because the expression of HB-EGF in asthmatic lungs was significantly elevated, we can speculate that progenitor cells that can further differentiate into smooth muscle–like cells might be recruited to lung tissue by HB-EGF, and at least part of them may play a role in the ASM layer thickening.

In conclusion, we have provided evidence that HB-EGF can stimulate ASM cell migration, and p38 MAPK-dependent actin cytoskeleton remodeling is involved in cell migration. Meanwhile, HB-EGF–induced smooth muscle progenitor cell migration from circulation to asthmatic lung might be another source for ASM remodeling in asthma.

We thank the staff members of this trial, colleagues, and all of the study staff for the enormous efforts in collecting and ensuring the accuracy and completeness of all of the data.

The authors have no financial conflicts of interest.