Faculté de Médecine, Faculté des Sciences Université Paris XII, Institut National de la Santé et de la Recherche Médicale, Unité 492 de Physiopathologie et Thérapeutique Respiratoire, 94010 Créteil; Faculté de Médecine, Laboratoire d'Enzymologie et de Chimie des Protéines, 37032 Tours Cedex; and Département des Sciences du Vivant, Service de Neurovirologie, Commissariat à l'Énergie Atomique, 92265 Fontenay aux Roses, France
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ABSTRACT |
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Epidemiological and experimental
studies suggest that diesel exhaust particles (DEPs) may be associated
with increased respiratory mortality and morbidity. Several recent
studies have also shown that DEPs increase the production of
inflammatory cytokines by human bronchial epithelium (HBE) cells in
vitro. The present study investigates the effects of DEPs on the
interaction of l-HBE cells (16HBE14o-) with the cell and matrix
microenvironment based on evaluation of integrin-type cell/matrix
ligand expression, cytoskeleton (CSK) stiffness, and matrix remodeling
via matrix metalloproteinase (MMP)-1, MMP-2, and MMP-9 expression. The
results showed that DEP exposure induced: 1) a net
dose-dependent decrease in CSK stiffness through actin fibers,
2) a concomitant specific reduction of both
3- and
1-integrin subunits extensively
expressed on the HBE cell surface, 3) a decrease in the
level of CD44, which is a major HBE cell-cell and HBE cell-matrix
adhesion molecule; and 4) an isolated decrease in MMP-1
expression without any change in tissue inhibitor of matrix
metalloproteinase (TIMP)-1 or TIMP-2 tissue inhibitors. Restrictive
modulation of cell-matrix interaction, cell-cell connection, CSK
stiffness, and fibrillary collagen remodeling results in a decreased
wound closure capacity and an increased deadhesion capacity. In
conclusion, on the basis of these results, we can propose that, in
addition to their ability to increase the production of inflammatory
cytokines, DEPs could also alter the links between actin CSK and the
extracellular matrix, suggesting that they might facilitate HBE cell
detachment in vivo.
diesel exhaust particle
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INTRODUCTION |
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A PROGRESSIVE CHANGE IN air pollution has been observed over the last 15 years, characterized by high concentrations of atmospheric hydrocarbons, nitrogen oxides, ozone, and especially respirable particulate matter (PM 10, PM 2.5, and now PM 0.1; see Ref. 16). These particles are mainly derived from diesel combustion, which is particularly high in France because of the extensive development of diesel engines (30% of all cars in France in 2000; see Ref. 38). Diesel exhaust particles (DEPs) are composed of a carbonaceous core with adsorbed traces of heavy metals and a vast number of organic compounds, such as polyaromatic hydrocarbons (45). DEPs tend to form aggregates 0.1-0.5 µm in diameter, placing these particles within the respirable range (20).
Airway epithelial cells are the primary target for air pollution and
may play a key role in the pathophysiology of airway diseases. Recent
studies have emphasized the role of DEPs in the development of an
inflammatory response of bronchial epithelial cells. In vitro studies
have shown that exposure of human bronchial epithelial (HBE) cells to
DEPs significantly increased cell electrical resistance, decreased
ciliary beat frequency (4), and induced release of
interleukin (IL)-1, IL-8, and granulocyte-macrophage colony-stimulating factor (GM-CSF) inflammatory cytokines (4, 6,
7). However, although the proinflammatory impact of DEPs on
bronchial epithelium is starting to be elucidated, little or no
information about the other possible biological effects of DEPs is
available at the present time. In particular, bronchial epithelial
cells establish intercellular tight junctions and adhere to underlying
basement membranes, which in turn may regulate bronchial epithelial
cell shape, cell proliferation, and differentiation. In pulmonary
diseases such as asthma, damage to bronchial epithelium is often
associated with disruption of the underlying basement membrane and
cell-cell interactions (34). On the basis of these data,
it appeared important to determine whether DEPs may impair the
interaction of bronchial epithelial cells with the matrix and cellular microenvironment.
The present study investigates modulation of cell-cell interactions between bronchial epithelial cells in response to DEP exposure, in terms of E-cadherin and CD44 (Cluster of Differentiation) adhesion molecule expression, since E-cadherin is responsible for homotopic adhesive interactions with E-cadherin on the surface of opposing cells (19), whereas hyaluronic acid receptor CD44 has a variety of functions, including participation in cell-cell adhesion by heterotopic and homotopic adhesions as well as cell-matrix interactions (11, 36).
Cell-matrix interactions between bronchial epithelial cells and
underlying basement membrane were also investigated by evaluating integrin expression in response to DEPs. Each transmembrane integrin is
composed of - and
-subunit heterodimers, and the combination of a
particular
-subunit with a given
-subunit imparts a defined matrix ligand specificity to each integrin (8, 23). In
addition, because integrins link the extracellular matrix (ECM)
components to the cytoskeleton (CSK) via actin filaments (F-actin), and
since the focal adhesion points mediated by these integrins are
involved in cell shape, stiffness, and migration, actin CSK stiffness
in response to DEP exposure was also investigated.
Adhesive contacts may be broken by extracellular proteolytic matrix metalloproteinases (MMPs). MMPs degrade all components of the ECM and play key roles in normal physiological processes involving ECM remodeling, such as wound healing, angiogenesis, or development. More particularly, MMP-1 cleaves fibrillary collagen at a unique site in the triple helix of the protein, whereas MMP-2 and MMP-9 (gelatinases A and B, respectively) specifically degrade gelatin and type IV collagen, and to a lesser extent, laminin, entactin, fibronectin, fibrillin, and elastin. Our previous studies reported that these two gelatinases were expressed by HBE cells (9, 30, 46). MMP activity is also regulated by specific tissue inhibitors of matrix metalloproteinases (TIMPs; see Ref. 35), and our previous studies have demonstrated that the two tissue inhibitors (TIMP-1 and TIMP-2) were also expressed by HBE cells (47, 48). The balance between these MMPs and TIMPs present in these cells in response to DEP exposure was therefore examined.
Overall, our results consistently showed that DEPs are able to alter bronchial epithelial cell-matrix interactions and epithelial cell-cell connections and ECM remodeling, in association with increased deadhesion cell capacity and a decreased rate of cell repair in response to mechanical injury.
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MATERIALS AND METHODS |
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Cell Cultures
All parameters were investigated in cultures of a HBE cell line (16HBE14o-), allowing us to perform a large number of reproducible experiments by using methods as different as flow cytometry, magnetic twisting cytometry (MTC), or gel zymography. Some primary HBE cell cultures were also carried out to compare the data obtained with the 16HBE14o- cell line.Isolation and culture of primary HBE cells. HBE biopsies were obtained by fibroscopy in several patients investigated for lung cancer. Biopsies were taken away from the tumor. Pathological examination confirmed the presence of normal bronchial mucosa on each specimen. All the patients whose biopsies were used for collection of epithelial cells gave their informed written consent, and the experimental design was approved by a relevant ethical committee (Comité Consultatif pour la Protection des Personnes dans la Recherche Médicale) in accordance with good clinical practice and French bioethical laws.
Primary (p) HBE cells were cultured according to the modified method of Baeza-Squiban et al. (3). Two or three explants (0.5 × 0.5 mm) were placed on sterile 35-mm plastic dishes (Nunc, Napperville, IL) coated with a collagen G matrix (bovine type I and III collagens; Biochrom, Berlin, Germany). The explants were covered with 600 µl of culture medium and incubated for 24 h. Culture medium (2 ml) was then added to each dish. The culture medium consisted of serum-free DMEM-Ham's F-12 (1:1; Life Technologies, Cergy-Pontoise, France) supplemented with 2% Ultroser G (UG; Life Technologies), 1% antibiotics (10,000 U/ml penicillin G sodium, 10,000 µg/ml streptomycin sulfate), 25 µg/ml amphotericin B (Life Technologies), and 2 mM glutamine (Life Technologies) at a final concentration. Explants were placed in a humidified incubator at 37°C in 5% CO2-95% air. The culture medium was changed every 2 days. Cells were cultured for 2 wk until confluence of p-HBE cells. The same explants were also successively transferred to new sterile coated plastic dishes, at 5- to 8-day intervals, to initiate new p-HBE cell cultures.HBE cell line culture. The 16HBE14o- cell line (a gift from Dr. D. C. Gruenert, University of Vermont, to B. Housset, Centre Hospitalo-Universitaire Henri Mondor, Créteil, France) was cultured on collagen G- and matrix-coated dishes. Different dish types (Nunc) were used depending on the experiment: 48-well plates for cytotoxicity assays, 24-well plates for wound healing assays, 8 Lab-Tek wells for actin structure investigation and for proliferating cell localization during wound healing, 32-mm2 dishes for MTC, and 25-cm2 flasks for flow cytometry assays. The culture medium was the same as that described above. Experiments were performed at passages 25 to 40, and all cultures were incubated in a humidified incubator at 37°C in 5% CO2-95% air. The culture medium was changed every 2 days.
Cell treatment by DEPs or carbon black. Carbon black, which represents the carbonaceous core of DEPs, was obtained from Sigma (L'Ile d'Abeau Chêne, France) and was used to test the involvement of the carboneous core in comparison with entire DEPs. Diesel PM SRM 1650 was purchased from the National Institute of Standards and Technology (Gaithersburg, MD). Stock solutions of particles were dispersed in 0.04% dipalmitoyl phosphatidylcholine (DPC; Sigma) in distilled water and then sonicated in ice for 5 min at maximum power (8 kilocycles). As previously described (6), DPC was used to mimic the lung surfactant lipid, and entire DEPs or carbon black were used at the final concentration of 1-100 µg/ml in the cell culture medium. Equivalent volumes of DPC alone were also used as DPC controls for each particle concentration. Subconfluent cultures of l-HBE cells were treated in the presence of particles for 24 h and compared with nontreated cells (controls) or DPC-treated cells. Concerning p-HBE cells, they were first incubated in culture medium for 24 h (control cells) and then treated with DPC solvent, DEPs, or carbon black for the next 24 h (treated cells) after changing the culture medium.
Cytotoxicity Assays
DEP, carbon black, or DPC cytotoxicity was evaluated using Neutral Red vital staining by estimating the cell viability measured as the uptake of the dye. Nontreated l-HBE cells or cells treated with DPC or particles were washed two times in PBS and were incubated for 1 h in the presence of 10 µg/ml Neutral Red (Sigma). After removing the culture medium and three washes with PBS, Neutral Red associated with living cultured cells was extracted with 200 µl/well 1% acetic acid in 50% ethanol, atDEP, carbon black, or DPC cytotoxicity after 24 h of treatment was also evaluated using the methylthiazoletetrazolium (MTT) assay. Briefly, exponentially growing cell culture cells were incubated for 90 min in the presence of 500 µg/ml MTT (Sigma). After removing the culture medium, reduced MTT product associated with viable cells was extracted with 400 µl/well isopropanol containing 8% 1 N HCl, and absorbance was measured at 450 nm with a microplate photometer. Cytotoxicity measurements were carried out after 24 h of treatment. DMSO (3 and 4%) cytotoxicity was measured as a positive control.
Evaluation of Particle Phagocytosis or Cell Contact
Phagocytosis or cell contact of DEPs and carbon black was investigated by optical microscopy and flow cytometry.Optical microscopy. Treated l-HBE cells were dissociated with enzyme-free cell dissociation buffer (131510-14; GIBCO-BRL, Cergy-Pontoise, France). For each particle concentration, phagocytic cells were counted using a hemocytometer (Malassez device) with a optical microscope (Zeiss).
Flow cytometry. DEP, carbon black phagocytosis, or cell contact was investigated by flow cytometry (FACS LSR; Becton-Dickinson) after cell dissociation with enzyme-free cell dissociation buffer. The following two parameters were measured: 1) the side scatter (SSC) parameter, which assesses the more granular appearance of epithelial cells in response to phagocytosed or membrane-associated carbon black and DEPs and 2) the natural fluorescence of cells, which was enhanced only in response to phagocytosed and cell membrane-associated DEPs because of the autofluorescence of polyaromatic hydrocarbons adsorbed on the DEP surface (530 nm specific emission wave length for 488 nm specific excitation wave length). This property was used previously by other authors to investigate the polycyclic aromatic hydrocarbons associated with air pollutants (24, 42).
F-actin Staining with Fluorescent Phallotoxin and Confocal Microscopy
Control or treated l-HBE cells were rinsed two times with warm CSK buffer (in mM: 50 NaCl, 300 sucrose, 3 MgCl2, and 10 PIPES, pH 6.8), which maintains CSK integrity, as previously described (10). Cells were then fixed with CSK buffer supplemented with 1% glutaraldehyde (Sigma) for 15 min. Glutaraldehyde was removed, and samples were rinsed two times for 2 min with warm CSK buffer. Cells were permeabilized for 3 min with CSK buffer supplemented with saponin (25 µg/ml; Sigma) and were then rinsed three times with CSK buffer. Rhodaminated phalloidin (1.5 µM; Sigma) was dissolved in CSK buffer and added to each sample for 30 min in darkness and in a humid chamber at room temperature. Coverslips were rinsed two times for 5 min with CSK buffer, and the last rinse was performed with ddH2O. Coverslips were mounted on slides with the cell side facing downward in Vectashield mounting medium (Vector Laboratories). Samples were stored overnight at 4°C before examination by laser confocal microscopy using an LSM 410 inverted microscope (Zeiss), comprising two internal helium-neon lasers and one external argon ion laser. Image processing was performed using LSM 410 software. Cell fields were randomly selected, brought into focus using a ×40/1.25 or 63/1.25 numeric aperture Plan Neofluor objective under transmitted light bright-field conditions, and briefly examined. Cross-sectional images were recorded under confocal conditions and used to establish a plane of focus above the glass surface. Optical sections were recorded every 1 µm to reveal intracellular fluorescence, and image reconstitution was performed.CSK and Actin CSK Stiffness Evaluation by MTC
Control or treated l-HBE cells were incubated in UG-free medium supplemented with 1% BSA (Sigma) for 30 min at 37°C to block nonspecific binding. Carboxyl ferromagnetic beads (4.0-4.5 µm diameter; Spherotec) were coated with arginine-glycine-aspartic acid (RGD) peptide according to the manufacturer's procedure (Telios Pharmaceuticals). Coated beads were incubated in serum-free medium supplemented with 1% BSA for 30 min before use. Beads were then incubated with the cells (30 µg/dish) for 30 min at 37°C in 5% CO2-95% air, and unbound beads were washed away with serum-free medium supplemented with 1% BSA. Dishes were then placed in the magnetocytometer. A brief 1,500-G magnetic pulse was first applied to magnetize all surface-bound beads in a unique horizontal direction, and a magnetic torque was then generated by applying an orthogonal uniform magnetic field (42 G). Associated changes in angular strain of the beads were measured by an on-line magnetometer and subsequently transformed to analog data with an acquisition system (Acqknowledge III; BIOPAC). Stiffness was defined as the ratio of stress to angular strain (43). This ratio measures the cell capacity to resist a local deformation, corresponding to CSK stiffness (Pa). After a first measurement of CSK stiffness, cytochalasin D (cyto D; Sigma), an agent that depolymerizes actin microfilaments, was added (10 µg/ml) for 25 min, and CSK stiffness was measured again. The difference of CSK stiffness before and after cyto D was calculated (Pa) and was representative of actin CSK stiffness or cell tone.Quantification of HBE Cell Adhesion Molecule Level by Flow Cytometry Assays
Expression of cell surface adhesion molecules.
All washings were performed in PBS supplemented with 0.01%
NaN3 (Sigma). All centrifugations (500 g) were
performed for 10 min at 4°C. l-HBE or p-HBE cells were dissociated by
enzyme-free cell dissociation buffer (GIBCO-BRL). The cell suspension
was centrifuged and incubated for 30 min at 4°C in PBS supplemented with 0.01% NaN3 and 10% AB human serum (AbCys, Paris,
France) to avoid nonspecific binding to Fc fragments. Cells
(105 to 2 × 105) were then incubated with
saturating concentrations of either adhesion molecule-specific
antibodies, purified mouse or rat isotype-matched controls
(IgG1, IgG2a, or IgG2b; Immunotech,
Marseille, France), or preimmune goat serum (Life Technologies) for 30 min at 4°C. We used monoclonal antibodies for L,
M,
x,
1,
2,
3,
4,
5,
6,
v,
1,
2,
3,
CD44, E-cadherin antigens (Immunotech), or for
4
(Novocastra Laboratories, Newcastle, UK) and polyclonal antibodies for
5- and
6-antigens (Santa Cruz
Biotechnology). After incubation with specific antibodies or
isotype-matched controls, cells were washed two times with PBS
supplemented with 0.01% NaN3 and labeled with 50 µl
appropriate secondary antibody for 30 min [either FITC-conjugated
F(ab')2 rabbit anti-mouse immunoglobulins (DakoPatts,
Glostrup, Denmark), FITC-conjugated F(ab')2 goat anti-rat immunoglobulins, or FITC-conjugated F(ab')2 rabbit
anti-goat immunoglobulins (Jackson Immunoresearch Laboratories)].
These secondary antibodies were diluted in PBS supplemented with 0.01%
NaN3 and 10% FCS at dilutions of 1:20 for the first
antibody and 1:50 for the other two antibodies. Cells were washed two
times, fixed in 200 µl CellFix (Becton-Dickinson, San Jose, CA)
solution containing 1% formaldehyde and 0.1% NaN3, and
stored at 4°C in the dark until analysis. Each mean adhesion
molecule-specific fluorescence intensity (MFI) was determined for
104 viable cells selected from total cells using an LSR
flow cytometer (BD Bio Science) and was corrected for corresponding
isotype control MFI. To determine whether the action of DEP on adhesion
molecule expression was partly phagocytosis dependent, adhesion
molecule expression was also assessed in cells treated by cyto D (10 µg/ml) for 24 h in the presence of DEP (100 µg/ml).
Quantification of the adhesion molecule intracellular pool. For each experimental condition, cell surface adhesion molecules were immunostained as described above. Cells were then fixed and permeabilized with IntraPrep Permeabilization reagent according to the manufacturer's recommendations (Immunotech). Briefly, cells were fixed for 15 min at room temperature with 5.6% formaldehyde, washed with PBS, centrifuged for 5 min at 500 g, and gently resuspended with a saponin solution kit for 5 min. After cell permeabilization, intracellular adhesion molecules were then stained. Cells were washed two times, fixed in 200 µl CellFix solution, and stored at 4°C in the dark for <24 h. To quantify both membrane and intracellular adhesion molecules (total pool), the following four immunostaining combinations were performed: 1) cell membrane-specific adhesion molecule immunostaining followed by isotype-matched control immunostaining after cell permeabilization; 2) cell membrane isotype-matched control immunostaining followed by specific adhesion molecule immunostaining after cell permeabilization; 3) specific adhesion molecule immunostaining, before and after cell permeabilization; and 4) isotype-matched control immunostaining, before and after cell permeabilization. The intracellular pool of each adhesion molecule was calculated by substracting the cell-surface adhesion molecule pool from the total pool.
Gel Zymography
Culture supernatants were centrifuged, and samples were frozen atELISA assays
The levels of MMP-1, TIMP-1, and TIMP-2 released in cell culture media were measured by human MMP-1, TIMP-1, and TIMP-2 ELISA kits (Amersham), according to the manufacturer's recommendations.Cell Deadhesion
After reaching confluency, l-HBE cells were dissociated with enzyme-free dissociation buffer, and the number of detached cells was counted with an hemocytometer (Malassez device) every 5 min for 40 min.In Vitro Wound Healing Assay
The in vitro wound healing assay was carried out as previously described (32, 37). A linear wound was made by gently scratching a subconfluent culture of l-HBE cells with a pipette tip followed by extensive rinsing with culture medium to remove all cellular debris. The area of denuded surface was quantified for 24 h. The plate was placed on an inverted microscope (Zeiss, Rueil-Malmaison, France), and an image was obtained by using a charge-coupled device camera (CCD-IRIS, Sony) connected to the microscope. The image was subsequently captured by an image-analyzing frame-grabber (Video enhancer FA320; FUTEK) and analyzed by image analysis software (Scion Image 1.3b). Wound closure was evaluated as the average surface covered by the cells after 3.5, 7, and 24 h of incubation and expressed as a percentage of exposed surface compared with the covered surface in the DPC control.Assessment of Cell Proliferation by Bromodeoxyuridine Incorporation
Localization of cell proliferation. l-HBE cells were plated on Lab-Tek 8 wells (Nunc) dedicated for immunofluorescence assay. At subconfluence, cells were either treated or not treated by particles or DPC for 24 h. A linear wound was then made as described above. Cultures were incubated for another 7 h at 37°C in humidified incubators under 5% CO2 in air, and 5-bromo-2'-deoxyuridine (BrDU) incorporation and detection were performed as previously described (17) using a BrDU Labeling and Detection Kit according to the manufacturer's recommendations (Boehringer-Mannheim). Samples were stored at 4°C overnight, and localization of cell proliferation was analyzed by laser confocal microscopy using an LSM 410 inverted microscope.
Quantification of cell proliferation by flow cytometry. Subconfluent l-HBE cells were incubated for 30 min at 37°C with the diluted (1:1,000) kit solution containing BrDU. Cells were then dissociated by enzyme-free cell dissociation buffer and were fixed and labeled as described above. Cells were stored at 4°C in the dark until analysis. BrDU specific fluorescence was quantified using an LSR flow cytometer.
Statistical Analysis
Statistical analysis was performed using nonparametric tests (Statview 5.0; SAS Institute). The Kruskal-Wallis test was used for multiple comparisons. Further comparisons were performed using Dunn's test. Significance was considered for P < 0.05. The significance of flow cytometry data was assessed by the Kolmogorov-Smirnov test, using CellQuest 3.1 software (Becton-Dickinson). ![]() |
RESULTS |
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Cytotoxicity of DEPs
DEPs, carbon black, and DPC cytotoxicity was evaluated on the basis of l-HBE cell staining by Neutral Red dye after 24, 48, and 72 h of treatment. A net significant cytotoxicity was found for the highest concentration of DEPs (100 µg/ml) and was time dependent since 31 ± 4 and 83 ± 8% of cells had died at 48 and 72 h, respectively. Lower levels of cytotoxicity were observed with carbon black (100 µg/ml) or equivalent DPC exposure for 48 and 72 h. Indeed, significant toxicity was observed in 20 ± 4 and 36 ± 5% of cells in response to 72 h of exposure to carbon black and DPC, respectively. The reliability of the method was attested by the high level of cytotoxicity (98%) induced by 5% DMSO at all times and by the lack of staining interference with collagen G matrix alone. No cytotoxicity was found after 24 h of treatment, whatever the treatment and the dose. This absence of any cytotoxicity was corroborated after 24 h of treatment, using MTT assay (data not shown). Because our results for longer culture times showed a time- and dose-dependent cytotoxicity induced by DEP exposure, as previously described (6, 40), subsequent investigations of DEP effects were evaluated on l-HBE cells cultured for 24 h.DEPs Are in Contact with the Cell Membrane and May Be Phagocytosed by HBE Cells
The number of l-HBE cells that were in contact with DEPs and carbon black particles at different concentrations (5, 20, and 100 µg/ml) and/or that have phagocytosed these particles was approximately evaluated by optical microscopy by counting the cells with an hemocytometer (Malassez device). Carbon black and DEP cell phagocytosis and/or cell contact was a dose-dependent process, and DEPs appeared to be more intensely in contact with the cell membrane and more phagocytosed than carbon black (Table 1).
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Flow cytometry assays were also performed for quantitative
determination of DEP phagocytosis and/or DEP cell contact. The DPC
solution used to disperse the particles had no effect on the SSC
parameter compared with control l-HBE cells. DEPs or carbon black, at
the concentration of 5 µg/ml, did not affect SSC, probably because of
the limit of sensitivity of flow cytometry. However, a significant net
shift of SSC was induced in response to 100 µg/ml DEP exposure (Fig.
1A), whereas a less marked
shift was observed in response to 20 µg/ml DEP exposure (data not
shown). Also, only DEP cell phagocytosis or DEP cell contact was able to induce a dose-dependent increase in l-HBE cell autofluorescence evaluated by flow cytometry (Fig. 1B).
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DEPs Are Engulfed by F-actin
Localization and distribution of F-actin staining were observed by confocal microscopy by performing optical cell sections every 1 µm from the apical to basal cell surface and after image reconstruction (Fig. 2). In the middle section of the confluent control and DPC-treated l-HBE cells (Fig. 2, A and B, respectively), intense and dense actin staining was observed at the cell periphery, whereas weak and scattered staining was observed in the cytosol. In carbon black-exposed l-HBE cells, internalized particles were clearly observed in the absence of any fluorescence (Fig. 2Cr), whereas little or no organized F-actin staining was observed around these internalized particles on adjacent serial sections (Fig. 2Cl). In DEP-exposed l-HBE cells, phagocytosed particles and particles located close to the cell membrane were observed in the absence of fluorescence (Fig. 2Dr), whereas colocalized and well-organized F-actin staining was observed on adjacent serial sections and generally engulfed these particles (Fig. 2Dl). Figure 2E presents a higher-power view of F-actin engulfing DEPs.
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DEP Exposure Induces a Dose-Dependent Decrease in Actin CSK Stiffness
Results for CSK stiffness and actin CSK stiffness (corresponding to the difference between CSK stiffness before and after cyto D) are shown in Fig. 3. The presence of DPC in the culture medium did not affect CSK stiffness or actin CSK stiffness (data not shown) of l-HBE cells. In contrast, DEP cell exposure induced a similar significant decrease in CSK stiffness, i.e., 24.4, 21, and 20% for the three levels of DEP concentration used (5, 20, and 100 µg/ml) and a dose-dependent decrease in actin CSK stiffness (34.4, 50, and 55%, respectively). No change in CSK stiffness or actin CSK stiffness was detected in response to DEP exposure at the lower concentration of 1 µg/ml (data not shown), confirming the dose-dependent response. Effects induced by carbon black exposure were usually associated with no change or a nonsignificant decrease in CSK stiffness and actin CSK stiffness (data not shown).
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DEPs Induce a Decrease in 3 (CD49)- and
1 (CD29)-Integrin Subunits and CD44 Receptor Expression
at the l-HBE Cell Surface
Expression of adhesion molecules at the l-HBE cell surface was never
modified by DPC. At the concentrations of 20 and 100 µg/ml, DEP
exposure induced a reproducible and significant dose-dependent decrease
in the expression of the three major adhesion molecules (3- and
1-integrin subunits and CD44
receptor). Figure 4 shows the shift
related to
3-,
1-, and CD44
immunofluorescence induced by the highest dose of DEPs. The apparent
increase in staining with the isotype control antibodies did not
reflect an increase in isotype control antibody binding but was only
because of the poly-aromatic hydrocarbons-induced autofluorescence of
l-HBE cells treated with DEPs. No effect of DEP exposure was
detected at the concentrations of 5 and 1 µg/ml (data not shown). The
effects induced by carbon black exposure were usually associated with no change or a nonsignificant decrease in
3- and
1-integrin subunits and CD44 receptor (data not shown).
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DEPs Induce a Decrease in Intracellular Distribution of
3- and
1-Integrin Subunits and CD44
Expression
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DEP Phagocytosis Is Not Required to Induce a Reduction of
3-,
1, and CD44 Expression at the HBE
Cell Surface
The results summarized in Fig. 5
show that 1) cyto D treatment alone induced a reproducible
and significant increase in the expression of 1,
3, and CD44 adhesion molecules, by 18, 26, and 39%,
respectively; 2) cyto D treatment in the presence of DEP did
not prevent the decreased expression of
1,
3, and CD44 adhesion molecules; and 3) the
decreased expression of
1,
3, and CD44
adhesion molecule in the presence of cyto D (39, 47, and 37%,
respectively) was therefore clearly higher than the decrease observed
in the absence of cyto D (29, 35, and 22%, respectively).
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DEP Exposure Induces a Specific Decrease in MMP-1 Protein Expression with No Change in TIMP-1 and TIMP-2 Expression by p-HBE and l-HBE Cells
To assess whether DEP exposure was associated with a change in ECM remodeling, the balance of MMPs/TIMPs was investigated. MMP-2, MMP-9, MMP-1, TIMP-1, and TIMP-2 protein expression was investigated in p-HBE and l-HBE cell supernatants.Gelatin gel zymography results clearly showed that MMP-2 and MMP-9
gelatinase protein expression, released for 24 h in p-HBE and
l-HBE cell supernatants, in proactivated and activated forms (72 and 68 kDa for MMP-2 and 92 and 88 kDa for MMP-9), was unchanged in response
to DPC, DEP, or carbon black exposure, regardless of the dose and
regardless of the type of cell culture. Figure 6 shows gel zymography results obtained
for the highest dose of particles. Concomitant TIMP-1 and TIMP-2
expression evaluated by ELISA assays also remained constant (Fig.
7). In contrast, exposure to the highest
DEP dose (100 µg/ml) induced a significant net decrease in MMP-1
protein expression and a discrete nonsignificant decrease at the lower
DEP dose (20 µg/ml), whereas DPC or carbon black exposure did not
alter this expression (Fig. 7). However, this MMP-1 decrease was only
observed in p-HBE cells.
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DEP Exposure Is Associated with an Increase in l-HBE Cell Deadhesion Capacity
HBE cells were left untreated or were treated with DPC, carbon black, or DEP for 24 h and then incubated for 40 min in enzyme-free cell dissociation buffer. Kinetic evaluation of cell detachment was performed every 5 min for 40 min. The results presented in Fig. 8 indicate that DEP exposure induced a dose-dependent amplification of cell detachment at the early times, since, after 5 min of incubation, 4% of l-HBE cells were still detached in response to 5 µg/ml DEP, whereas only 0.5% was detached in response to an equivalent dose of DPC. After 10 min of incubation, 60% of cells were detached, whereas only 22% of cells were detached in the presence of DPC alone. For a concentration of 20 µg/ml DEPs, the percentage of detached cells was 3% at 5 min and 82% at 10 min vs. 0.8 and 32%, respectively in the presence of DPC alone. Previous exposure to carbon black induced no change or a nonsignificant increase in the percentage of detached cells (data not shown).
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DEP Exposure Reduces l-HBE Cell Wound Repair Capacity
Kinetic studies of cell wound repair capacity were performed 3.5, 7, and 24 h after a linear mechanical wound performed on l-HBE cell confluent cultures. Wound closure, expressed as the average percentage of surface exposed compared with the surface covered in DPC controls, was not significantly changed in response to carbon black exposure. In contrast, DEPs inhibited wound closure in a dose-dependent fashion (Fig. 9). At 24 h, the wound was totally closed regardless of the type of treatment.
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Alteration of l-HBE Cell Repair Capacity Induced by DEP Exposure Is Partially Due to Decreased Cell Proliferation
At 7 h postwound, l-HBE cell proliferation was localized by immunofluorescence (BrDU incorporation and detection) and was similar at the edge of the wound and any other site of the cell culture (data not shown). To more precisely quantify a possible discrete change in HBE cell proliferation in response to DEP exposure, BrDU-specific fluorescence was analyzed by flow cytometry on subconfluent l-HBE cells. A weak and nonsignificant increase in cell proliferation was observed in response to DPC volume used to disperse DEPs at a concentration
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DISCUSSION |
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The main objective of the present study was to determine whether DEP exposure could alter the balanced interaction between HBE cells and their matrix microenvironment. Our data clearly demonstrate that DEP exposure induces a dose-dependent reduction of l-HBE cell actin CSK stiffness and matrix adhesion molecule expression, associated with impaired wound repair capacities and increased cell deadhesion potential. These results therefore suggest that L-HBE cells exposed to DEP pollution might lose their capacity to interact with the environmental ECM, thereby accentuating their shedding susceptibility.
DEPs Are in Contact with the Cell Membrane and May Be Phagocytosed by l-HBE Cells
First, to more clearly define our cell culture model, we analyzed contacts between l-HBE cells and DEPs by using three different approaches. Our optical microscopy analysis (Table 1), SSC determination by flow cytometry (Fig. 1A), and PAH-induced autofluorescence determination by flow cytometry (Fig. 1B) showed that DEPs were in contact with the cell membrane and/or phagocytosed by l-HBE cells in a dose-dependent fashion. However, neither of the three approaches was able to clearly distinguish between cell-ingested or cell-bound particles; evaluation of phagocytosis was therefore only approximate. The obvious more intense phagocytosis of DEPs compared with carbon black may be because of their different size. The particle size of Sigma carbon black was slightly greater than that of DEPs, and it has been proposed that small particles are more readily phagocytosed by epithelial cells than larger particles (12, 13). Previous studies (22) have also reported phagocytosis of DEPs by cat bronchial epithelial cells in vivo.DEP phagocytosis is shown by the pattern of distribution of F-actin fluorescent phallotoxin staining in l-HBE-cells (Fig. 2). In control cells, cortical actin filaments are mainly organized into stress fibers involved in adhesion plaques at the basal surface and in circumferential belts, whereas scattered cytosolic actin represents small and slightly polymerized actin. In l-HBE cells exposed to DEPs, organized F-actin often engulfed the phagocytosed DEPs and also clearly extended as lamellipodia around the DEPs close to the cell membrane. This process appeared to be much more intense for DEPs than for carbon black particles and may be because of the presence of various organic compounds and metals adsorbed on the DEP surface.
DEP Exposure Induces Alteration of Cell-Matrix and Cell-Cell Interactions
In the present study, we used MTC to evaluate possible changes in HBE cell CSK stiffness and more particularly changes in actin CSK stiffness in response to DEP exposure. MTC is a relatively new technique used to measure CSK stiffness by application of controlled mechanical stress applied directly to cell-surface integrins, using RGD-coated microbeads (43). This type of CSK stiffness measurement is directly proportional to the strength of cell-matrix interactions (29, 44), and the difference between stiffness before and after brief exposure (25 min) to cyto D reflects actin CSK stiffness. Our results demonstrated that CSK stiffness of l-HBE cells was significantly reduced by 20% in response to DEP exposure and that actin CSK stiffness was reduced in a dose-dependent fashion by 35, 50, and 55% for 5, 20, and 100 µg/ml DEPs, respectively (Fig. 3). Compared with the variable effect of carbon black, these results suggest that DEPs are specifically able to induce loss of CSK stiffness.Also, DEPs are specifically able to induce a loss of interactive
cell-ECM properties, since flow cytometry quantification of the
expression of all integrins at the l-HBE cell surface showed a
significant dose-dependent decrease between 20 and 40% in major 3- and
1-integrin subunits and CD44
receptor in response to DEP exposure (Fig. 4). This decrease in both
3- and
1-integrin subunits strongly
suggests a concomitant decrease in
3
1-integrin heterodimer. The
3
1-integrins and CD44 hyaluronic acid
receptor have already been described in HBE cells (5, 33),
and the marked decrease in these two major molecules in response to DEP exposure indicates that they play a key role in the loss of l-HBE cell-matrix interactions. Together, these data strongly suggest that
the DEP-induced decrease in actin CSK stiffness may be at least partly
because of the DEP-induced loss of adhesion molecules, in addition to
the different F-actin distribution pattern. There is now increasing
evidence that integrins regulate CSK assembly via a signaling pathway.
More particularly, in hepatic stellate cells, Kojima et al.
(25) observed that cell surface integrin binding to
interstitial collagen induced a change in their CSK via a signal
transduction system. Nevertheless, some studies have shown that the CSK
itself may actively regulate integrin expression and binding
conformation as well their mobility in the membrane (2).
Moreover, because 3
1-integrins and
CD44 receptor are also known to be involved in cell-cell interactions
(28, 41), their downregulation induced by DEP exposure may
result in loss of cell-cell interconnections, predisposing to cell
detachment. Our results confirm this predisposition, since the
percentage of detached l-HBE cells in response to dissociation buffer
was higher in the presence of DEPs (Fig. 8). Finally, we propose that
this higher deadhesion capacity probably reflects an alteration of
structural links between CSK actin and the ECM via
3
1-integrin and CD44 downregulation.
Reduced expression of
3
1-integrins
induced by puromycin aminonucleoside was also recently proposed to
contribute to glomerular epithelial cell detachment from the glomerular
basement membrane (26).
Decrease in 1,
3, and CD44 Expression
by l-HBE Cells in Response to DEP Exposure Is Not Due to
Intracellular Internalization or Cytoplasmic Sequestration
Decrease in 1,
3, and CD44 Expression
by l-HBE Cells in Response to DEP Exposure Is Not Due to DEP
Phagocytosis
DEP Exposure Is Associated with a Reduction of the HBE Cell Repair Process
In agreement with our postulate that DEP exposure can alter the link between actin CSK and the ECM via a reduction in adhesion molecule expression, we also observed that, at 3.5 and 7 h, in vitro wound closure was dose dependent and decreased in the presence of DEP (Fig. 9) but seemed to occur without any apparent signs of cell migration or cell spreading. In general, cell locomotion is a dynamic interplay between cell-ECM adhesion, extension of the leading edge of the cell, and retraction of the trailing edge. To move along the ECM, cells, and more particularly p-HBE cells, first adhere to the matrix by establishing primordial cell-ECM contacts distributed at the cell front heading in the direction of migration (30, 39). In contrast with p-HBE and type II alveolar pneumocyte repair models (37), no migrating or spreading cell morphology was identified at the leading edge of the wound during the repair process of our l-HBE cell repair model. In the latter, cell proliferation appeared to be the key factor during wound healing and was decreased in response to DEP exposure, as stated by quantitative BrDU incorporation data (Fig. 10).Taken together, these results, demonstrating a reduction of the cell
repair process, decreased cell proliferation, and decreased 3
1-integrin and CD44 receptor expression,
are in accordance with previous studies indicating a possible
relationship between the level of integrin or CD44 expression and cell
functions, such as cell repair or cell proliferation. High expression
of CD44 was found during in vitro repair of bronchial epithelial cells after a mechanical injury (31). CD44 can also stimulate
cell proliferation (14), and CD44 upregulation has been
demonstrated in areas of damaged bronchial epithelium both in normal
and asthmatic subjects, strongly suggesting that CD44 may be directly
or indirectly associated with repair processes occurring after damage
(27). As for integrins, a new concept is emerging to
suggest that integrins are multifunctional and may contribute not only
to epithelial cell adhesion but also to regulation of cell growth and
proliferation via a signaling pathway involving mitogen-activated
protein kinase (18). More particularly, proliferation of
human epithelial cells was significantly inhibited by a
function-altering
3-integrin antibody (18).
DEP Exposure Appears to Weaken Matrix Remodeling by Interstitial Collagenase MMP-1 Expressed by l-HBE Cells
Our results concerning ECM remodeling by subconfluent l-HBE or p-HBE cells did not demonstrate any change in MMP-2 or MMP-9 protein expression (Fig. 6) or TIMP-1 and TIMP-2 levels (Fig. 7) in response to DEP exposure. This result emphasizes the persistence of a harmonious balance between matrix gelatinases and their tissue inhibitors. Because MMP-9 expressed by p-HBE cells during wound repair has been recently proposed to play a key role in remodeling primordial contacts via specific degradation of type IV collagen (9, 30), unchanged MMP-9 expression appears relevant to the absence of any migrating event associated with our cell model in response to DEP exposure. De-Bentzmann et al. (15) recently suggested that MMP-2 overactivation associated with a limited increase in TIMP-2 was responsible for inhibition of Pseudomonas aeruginosa wound closure in vitro. In contrast, our results suggest that DEP wound closure inhibition would not depend on MMP-2 activation, probably because of different proteolytic mechanisms. Interestingly, interstitial collagenase MMP-1 production by l-HBE cells appeared to be markedly decreased in response to the highest dose of DEPs (Fig. 7), supporting the hypothesis that DEP exposure may reduce interstitial matrix turnover. Conversely, but not in opposition to the present study, recent studies by our group have demonstrated that the addition of exogenous MMP-1 collagenase to cultured alveolar epithelial cells could enhance wound healing by promoting cell migration on type I collagen (37).In conclusion, the present study proposes that, besides inducing a
well-known inflammatory reaction, DEP exposure of l-HBE cells could
alter cell-matrix interactions and cell cohesion via a concomitant
decrease in 3
1-integrin subunits and CD44
adhesion molecule and a reduction of actin CSK stiffness. Alteration of cell-matrix interactions and cell cohesion results in a dose-dependent reduction of the wound closure capacity and enhancement of the cell
deadhesion capacity. A reduction in wound closure also appears to
result from reduced cell proliferation and probably from alteration of
matrix remodeling via an imbalance between MMP-1 and TIMP-1 and -2 in
favor of inhibitors. Overall, these results support the concept that
DEP exposure tends to break the link between actin CSK and the ECM,
suggesting an increased potential for cell detachment from the
underlying basement membrane in vivo. Finally, the fact that there was
nonsignificant or no effect of carbon black particle exposure on actin
CSK stiffness, cell adhesion molecule expression, wound repair
capacity, the cell deadhesion process, or cell proliferation supports a
key role for polycyclic aromatic hydrocarbons adsorbed on the surface
of DEP.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Bernard Maitre for providing human bronchial biopsies and Antoine Mary for assistance with magnetic twisting cytometry.
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FOOTNOTES |
---|
* B. Doornaert and V. Leblond contributed equally to this work.
This study was supported by Institut National de la Santé et de la Recherche Médicale, Primequal-Predit grant from Ministère de l'Aménagement du Territoire et de l'Environnement, and a grant from Société Total Fina Elf.
Address for reprint requests and other correspondence: C. Lafuma, INSERM U492 de Physiopathologie et Thérapeutique Respiratoires, Faculté de Médecine, 8 rue du Général. Sarrail, 94010 Créteil Cedex (E-mail: chantal.lafuma{at}creteil.inserm.fr).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
August 23, 2002;10.1152/ajplung.00039.2002
Received 23 January 2002; accepted in final form 9 August 2002.
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