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Published ahead of print on July 29, 2004, doi:10.1165/rcmb.2003-0432OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 31, pp. 517-527, 2004
© 2004 American Thoracic Society
DOI: 10.1165/rcmb.2003-0432OC

Differential Regulation of Human Lung Epithelial and Endothelial Barrier Function by Thrombin

Kamon Kawkitinarong, Laura Linz-McGillem, Konstantin G. Birukov and Joe G. N. Garcia

Division of Pulmonary and Critical Care Medicine, Johns Hopkins University School of Medicine, Baltimore, Maryland; and Department of Medicine, Chulalongkorn University, Bangkok, Thailand

Address correspondence to: Joe G. N. Garcia, M.D., Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Circle 4B-77, Baltimore, MD 21224-6801. E-mail: drgarcia{at}jhmi.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Lung epithelial and endothelial barrier dysfunction is critical to the physiologic derangement observed in acute lung injury, but remains poorly understood. We utilized human alveolar epithelial (A549) and endothelial cells (EC) to study cytoskeletal remodeling, myosin light chain (MLC) phosphorylation and barrier regulation evoked by the edemagenic agent, thrombin. Thrombin-challenged human EC monolayers demonstrated increased MLC phosphorylation, actin stress fiber formation and loss of barrier integrity reflected by decreased transmonolayer electrical resistance (TER). In contrast, thrombin produced prominent circumferential localization of actin fibers, increased MLC phosphorylation and increased TER across epithelial monolayers, consistent with barrier protection. Reductions in MLC phosphorylation induced by cell pretreatment with pharmacological inhibitors of MLC kinase (ML-7) and Rho kinase (Y-27632) significantly attenuated thrombin-mediated TER changes and MLC phosphorylation in both lung cell types. Thrombin-produced, time-dependent activation of Rho GTPase in both epithelial and EC, whereas Rac GTPase activation was observed only in A549 cells. Molecular inhibition of Rac activity by adenoviral transfer of dominant-negative Rac mutant abolished thrombin-induced TER increases in alveolar epithelial cells. Finally, A549 cells, but not endothelium, demonstrated increased levels of tight junction proteins (ZO-1 and occludin) after thrombin at the cell-cell interface areas linked to thrombin-elicited barrier protection. These results demonstrate differential pulmonary endothelial and alveolar epithelial barrier regulation via unique actomyosin remodeling and cytoskeletal interactions with tight junction complexes, which confer selective barrier responses to edemagenic stimuli.

Abbreviations: acute respiratory distress syndrome, ARDS • bovine serum albumin, BSA • dominant-negative, DN • endothelial cells, EC • fetal bovine serum, FBS • G-protein–binding domain, GBD • human bronchial epithelial cell, HBEC • hepatocyte growth factor, HGF • human pulmonary artery EC, HPAEC • horseradish peroxidase, HRP • immunoglobulin, Ig • myosin light chain, MLC • MLC kinase, MLCK • protease-activated receptor, PAR • phosphate-buffered saline, PBS • polyvinylidene fluoride, PVDF • diphosphorylated MLC, ppMLC • Rho-associated kinase, RhoK • sphingosine 1-phosphate, S1P • Tris-buffered saline, TBS • TBS–Tween 20, TBS-T • transmonolayer electrical resistance, TER • thrombin receptor–activating peptide, TRAP


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The gas exchanging air–blood barriers in the lung are composed of the microvascular pulmonary endothelium and the epithelial lining of the alveoli, which form three separate compartments: blood, interstitium, and the alveolar space. Acute lung injury and its most severe form, the acute respiratory distress syndrome (ARDS), are conditions characterized by a diffuse, intense inflammatory process, by damage to both endothelial and epithelial cell barriers, resulting in marked extravasation of vascular fluid. The filling of alveolar spaces by proteinaceous edema fluid and inflammatory cells leads to severe hypoxemia and respiratory failure.

Thrombin, a key blood coagulation serine protease generated from a circulating prothrombin precursor after tissue injury, converts fibrinogen to fibrin, resulting in microthrombi. In addition, thrombin binds G-protein–coupled protease-activated receptors (PAR-1, PAR-3, and PAR-4) and promotes numerous cellular effects, including chemotaxis, proliferation, extracellular matrix turnover, and release of specific inflammatory mediators, vasoactive substances, and growth factors involved in tissue repair, inflammatory, and fibroproliferative processes. Thrombin activation is a putative key event in acute lung injury given that microthrombi in the pulmonary microvasculature and fibrin deposition in intra-alveolar and interstitial compartments are hallmarks of the early stages of ARDS. Exposure of plasma components to tissue factor expressed by endothelial cells (EC), macrophages, alveolar epithelial cells, or fibroblasts leads to intra-alveolar activation of coagulation and thrombin generation (1). Recent studies have demonstrated that successful anticoagulation associated with the reduction of thrombin levels attenuated lung microscopic pathology in animal models of ARDS (2) and decreased the mortality rate in patients with sepsis. These observations further support the significance of coagulation abnormalities in the pathogenesis of ARDS (1, 3) and sepsis, processes characterized by endothelial and epithelial barrier disruption.

Thrombin-induced EC barrier dysfunction and its regulatory pathways involve multiple signaling pathways and cytoskeletal targets, including Rho-kinase (RhoK) (4, 5) and the Ca2+/calmodulin-dependent myosin light chain (MLC) kinase (MLCK) (6–9). Increased RhoK and MLCK activation enhances actin–myosin interaction, central stress fiber formation, EC contraction, and gap formation indicative of barrier disruption and increased vascular permeability (10, 11). In addition, endothelial barrier regulation is not only dependent on the levels of MLC phosphorylation, but is also tightly linked to the spatial location of phosphorylated MLC (10). Whereas thrombin-induced activation of the small GTPase RhoA and RhoK can lead to direct MLC phosphorylation, RhoK-mediated phosphorylation and inactivation of the myosin-associated phosphatase, represents the primary mechanism of prolonged EC contraction and increased permeability.

In contrast to the well-described molecular mechanisms of thrombin-induced EC activation, the effect of thrombin on pulmonary epithelial cell barrier regulation is unknown. Despite extensive alveolar damage in the acute phase of ARDS, alveolar type 2 cells can survive and restore alveolar epithelium in the resolution phase of this syndrome (12). Thus, we hypothesized that the alveolar type 2 cells might have specific responses to external stimuli and examined the effects of thrombin on pulmonary epithelial cell barrier integrity, actomyosin cytoskeletal reorganization, and Rho and Rac-mediated signaling pathways. Because intercellular junctions, such as tight junctions and adherens junctions, are closely related to actin cytoskeleton–related barrier regulation in both epithelial cell and EC (13), we analyzed cell junction protein complex stability after thrombin stimulation. Our results indicate a novel, barrier-protective effect of thrombin on epithelial cells and suggest a critical role for cell-specific cytoskeletal remodeling and tight-junction regulation in endothelial and alveolar epithelial barrier regulation.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Reagents and Antibodies
Unless otherwise specified, reagents were obtained from Sigma (St. Louis, MO). Fetal bovine serum (FBS) was obtained from American Type Culture Collection (Manassas, VA). EC culture basal medium (EBM-2) with growth supplements and bronchial epithelial growth medium were obtained from Clonetics (Walkersville, MD). The following antibodies were commercially obtained: anti–ZO-1 and anti-occludin monoclonal antibodies (Zymed Laboratories, South San Francisco, CA), anti-ß tubulin antibody (Covance, Berkeley, CA), anti–diphospho-MLC–specific antibodies, which recognize phosphorylated serine19 and threonine18 of myosin regulatory light chains (Cell Signaling, Beverly, A), anti-ß catenin antibody (Transduction Laboratory, Lexington, KY), and anti–pan-MLC antibody. Specific peptide activator of protease-activated receptor-1 thrombin receptor–activating peptide (TRAP)-6 (SFLLRN) was obtained from Anaspec (San Jose, CA). The Rho activation assay kit (rabbit polyclonal anti-Rho antibody and glutathione S-transferase–tagged fusion protein corresponding to Rhotekin Rho-binding domain bound to glutathione-agarose), and the Rac activation assay kit (mouse monoclonal anti-Rac antibody and p21-activated kinase bound to agarose) were obtained from Upstate Biotechnology (Lake Placid, NY). Alexa Fluor 488 anti-mouse immunoglobulin (Ig) G antibody and Texas red-phalloidin were purchased from Molecular Probes (Eugene, OR). Horseradish peroxidase (HRP)–linked anti-mouse and anti-rabbit IgG antibodies and HRP Western blot detection kit were obtained from Cell Signaling (Beverly, MA). The protease inhibitor cocktail set, RhoK inhibitor Y-27632 and MLCK inhibitor ML-7 were obtained from Calbiochem (La Jolla, CA). Polyvinylidene fluoride (PVDF) membrane was purchased from Millipore (Bedford, MA). The recombinant adenovirus RacN17, containing the c-Myc–tagged dominant-negative (DN) form of Rac1, was kindly provided by Dr. Kwang Sik Kim (Johns Hopkins University, Baltimore, MD).

Human Pulmonary Artery EC, A549 Pulmonary Epithelial Cell, and Primary Human Bronchial Epithelial Cell Cultures
Human pulmonary artery EC (HPAEC) were obtained from Clonetics (Walkersville, MD) and used at passages 4–7. Cells were cultured in completed medium containing 10% fetal bovine serum (FBS), endothelial growth supplement and 1% of antibiotics and antifungal solution (penicillin, 10,000 U/ml; streptomycin, 10 µg/ml; and amphotericin B, 25 µg/ml). EC were incubated at 37°C in humidified atmosphere of 5% CO2 air incubator and grown to monolayers with typical cobblestone morphology. Human A549 cells (American Type Culture Collection, Manassas, VA) were cultured in Dulbecco's modified Eagle's medium, containing 10% FBS, antibiotics, and antifungal solution and used at passages 5–12. Primary human bronchial (HBEC), obtained from Dr. V. Natarajan (Johns Hopkins University, Baltimore, MD), were used at passage 1.

Measurement of Transendothelial and Transepithelial Cell Electrical Resistance
Endothelial and epithelial cells were grown to confluence in polycarbonate wells containing evaporated gold microelectrodes (surface area, 10–3 cm2) in series with a large gold counter electrode (1 cm2) connected to a phase-sensitive lock-in amplifier, as we have previously described (14). The size of the small gold electrode is critical for the impedance resulting from the presence of cells on the electrode to predominate over the resistance of the medium. Measurements of transmonolayer electrical resistance (TER) were performed using an electrical cell-substrate impedance sensing system (Applied BioPhysics Inc., NY), as we have described previously (14, 15). Briefly, current was applied across the electrodes by a 4,000-Hz AC voltage source with an amplitude of 1 V, in series with a 1 M resistance to approximate a constant current source (1 µA). The in-phase and out-of-phase voltages between the electrodes were monitored in real time with the lock-in amplifier and subsequently converted to scalar measurements of transmonolayer impedance, of which resistance was the primary focus. These methods have demonstrated a highly sensitive biophysical assay that indicates the state of cell shape and focal adhesion. The culture medium was replaced with free-serum basal media; TER was monitored for steady state, achieved to establish a baseline resistance. Agonist-mediated permeability was evaluated by measurement of TER. Values from each microelectrode were pooled at discrete time points and plotted against mean time ± SEM.

MLC Phosphorylation
Endothelial and epithelial A549 cells were grown to confluence in 35-mm dishes and stimulated with thrombin (50 nM) or vehicle, as indicated. The total lysates were collected and protein concentrations were measured by bicinchoninic acid (BCA) assay. Equal amounts of protein were used for all Western blot analyses. Activation of MLC phosphorylation was assessed by Western blot analysis with anti-diphosphorylated MLC (ppMLC) antibody, followed by membrane reprobing with anti-pan–MLC antibody, and quantitative scanning densitometry, as we have previously described (11). In selected experiments, the cells were preconditioned with either ML-7 (10 µM), or Y-27632 (5 µM) for 1 h before thrombin challenge.

Immunofluorescence Microscopy
Endothelial and epithelial cells grown on gelatinized cover slips were incubated with thrombin (50 nM) or vehicle control for predetermined times, washed with phosphate-buffered saline (PBS), fixed in 3.7% formaldehyde in PBS (15 min), permeabilized with 0.25% Triton X-100 in 0.1% Tris-buffered saline (TBS)–Tween 20 (TBS-T) (15 min), and blocked with 2% bovine serum albumin (BSA) in TBS-T (30 min). Incubations with primary antibody (anti–ZO-1, anti-occludin, anti–ß-catenin, and anti–ppMLC antibody) were performed in blocking solution (2% BSA in TBS-T) at 4°C overnight. After three washes with TBS-T, cells were incubated with the appropriate secondary antibody conjugated to immunofluorescent dyes (Alexa 488 for green fluorescence) in blocking solution and were kept in dark for 1 h at room temperature. For F-actin staining, Texas-Red–conjugated phalloidin (1:200) was used for 1 h at room temperature. After four washes with TBS-T, the cover slips were mounted. Analyses of immunofluorescent staining were performed using a Nikon Eclipse TE 300 microscope with 60x objective lens (Nikon, Melville, NY) and digital Spot camera (Diagnostic Instruments, Sterling Heights, MI).

Rac and Rho GTPase Activation Assay
Rac and Rho activity was assessed as previously described (14). The "pull-down" assay was used to detect active Rac and Rho by utilizing the G-protein–binding domain (GBD) of a downstream target of Rac or Rho that selectively binds the protein in its active (GTP-bound) form. Binding of the GBD by Rac or Rho is stable enough to allow isolation of the GBD–Rac/Rho-GTP complex by glutathione affinity beads, and the precipitated GTPase can then be quantified by Western blotting. Briefly, EC and A549 epithelial cells grown in 100-mm dishes and rendered quiescent in basal media containing 2% FBS overnight were incubated with 50 nM of thrombin in the same medium for 0, 5, and 30 min. After washing with PBS, cells were lysed in 500 µl of Mg2+ lysis buffer, containing 25 mM HEPES, pH 7.5, 150 mM NaCl, 1% NP-40, 10 mM MgCl2, 1 mM EDTA, 2% glycerol, 0.2 mM vanadate, 0.2 mM phenylmethylsulfonyl fluoride, and 1:200 dilution of phosphatase inhibitor cocktail, homogenized by pipetting, and then briefly centrifuged to remove the cell debris. For the Rac activation assay, supernatant (200 µl) was incubated with the agarose-conjugated p21-binding domain of human p21-activated kinase (residues 67–150; 10 µg) for 30 min. To determine the extent of Rho activation, supernatant (200 µl) was incubated with 20 µg of Rhotekin Rho-binding domain (residues 7–89) bound to glutathione-agarose for 45 min. The agarose beads were washed three times with Mg2+ lysis buffer and resuspended in 20 µl of 3x sodium dodecyl sulfate gel loading buffer. The samples were then subjected to electrophoresis in a 15% gel and transferred to PVDF membrane. Rho and Rac protein was detected by Western blot using rabbit polyclonal anti-Rho antibody and mouse monoclonal anti-Rac antibody, respectively. For total Rho and Rac detection, 5 µl of the original cell lysates were subjected to electrophoresis in the same manner.

Rac N17 Adenoviral Infection
The adenovirus encoding myc-tagged cDNA of DNRac1 mutant with T/N17 substitution was used for Rac downregulation in A549 cells. A549 grown on gold microelectrode plates for TER measurement were infected 48 h after plating with a multiplicity of infection of 10. Control cells were infected with adenovirus containing green fluorescent protein. After 24 h, the infection medium was aspirated and replaced with fresh complete medium. Adenoviral infection rate detected by immunofluorescence was ~ 95%. Expression of recombinant Rac protein was assessed by Western blotting using 10 µg of A549 protein lysates and a Rac-specific antibody.

Differential Protein Fractionation
HPAEC and A549 cells were seeded in D100 dishes and grown to 100% confluence. One hour after replacement of complete media by free-serum basal media, cells were treated with thrombin (50 nM) or vehicle control for 30 min. The monolayers of HPAEC and A549 from each dish were rinsed twice by PBS, and then were lysed in 500 µl extraction buffer (20 mM Tris-HCl, pH 7.4, 125 mM sucrose, 50 mM NaCl, 2 mM EGTA, 1 mM phenylmethylsulfonyl fluoride, 2 µl phosphatase inhibitor cocktails, and 2 µl protease inhibitor cocktails) on ice. The lysates were scraped, homogenized by passing several times through a 26-gauge needle, and collected in microtubes. After ultracentrifugation at 100,000 x g for 30 min at 4°C, supernatants containing cytosolic proteins and pellets containing particulate fractions were collected and resuspended in 3x sample buffer, then analyzed by Western blotting.

Western Blot Analysis of Cell Junction Proteins
Equal protein of total cell lysates and samples after differential fractionation were subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis. The separated proteins were electrically transferred to PVDF membranes. The blots were subsequently blocked with 5% BSA in TBS, containing 0.1% Tween 20 (TBS-T) at room temperature for 30 min and then incubated at 4°C overnight with primary antibodies (anti–ZO-1, anti-occludin, anti-ß catenin, or anti-tubulin antibody) for 1 h. After washing steps, membranes were incubated with the appropriate HRP-linked IgG secondary antibody at room temperature for 1 h. The protein bands of interest were then visualized with the enhanced chemiluminescence Western blot detection system according to the manufacturer's instructions. Blots were scanned and quantitatively analyzed using ImageQuant software (v5.2, Amersham Biosciences, NJ).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effect of Thrombin on TER and Actin Rearrangement
Thrombin (50 nM) produced a 40–60% decline in TER across HPAEC monolayers, reflecting increased permeability, as we have previously described (14, 16). This response was rapid, peaked after 15–20 min, and returned to baseline values by 4 h (Figure 1A). Similar thrombin stimulation of the human epithelial cell line (A549), however, induced rapid and significant increases in TER in a dose-dependent manner (Figure 1B). Importantly, the marked increase in TER in A549 cells was sustained and observed for up to 6–8 h at thrombin concentrations above 10 nM. These data suggest potentially distinct barrier-regulatory mechanisms exhibited by lung EC and epithelial cells after thrombin stimulation. Furthermore, we have extended these findings to other respiratory endothelial and epithelial cell types, including human lung microvascular EC and primary HBEC. Our results suggest that, similar to HPAEC, thrombin stimulation of human lung microvascular EC decreased TER, whereas thrombin stimulation of HBEC increased TER levels, consistent with the barrier-protective responses of A549 epithelial cells to thrombin challenge (Figure 1C).



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  		Figure 1. Differential effect of thrombin on lung epithelial and EC barrier properties. HPAEC and alveolar epithelial cells (A549) were plated on gold microelectrodes and grown to confluence. Electrical resistance across the EC (A) and epithelial cell (B) monolayers (TER) was measured as described in MATERIALS AND METHODS. Cells were stimulated with thrombin at the time marked by arrow. Thrombin induced significant TER decline in HPAEC monolayers, indicative of increased permeability (P < 0.05). In contrast, thrombin, in the range of concentrations from 10 nM to 500 nM, increased TER in A549 cultures in a dose-dependent manner, reflecting a barrier-protective response (P < 0.05). Transient thrombin-induced increases in TER in A549 were observed even at very low thrombin concentrations (1 nM) (data not shown). The barrier-protective effect of thrombin on lung epithelial cells was further confirmed using primary culture of bronchial epithelial cells and compared with thrombin effects on pulmonary macrovascular (HPAEC) and microvascular (HLMVEC) EC (C). TRAP-6, a PAR-1 agonist, increased TER similarly to thrombin response in A549 cells, indicating the role of PAR-1 receptor in thrombin-induced barrier protection in A549 (D). Shown are results of at least seven independent experiments. SE was < 0.03 ohms at each time point. (A: diamond, vehicle; square, thrombin (50 nM). B and C: diamond, Thr 0 nM; square, Thr 10 nM; triangle, Thr 100 nM; asterisk, Thr 500 nM. D: diamond, TRAP 0 µM; square, TRAP 1 µM; triangle, TRAP 10 µM; asterisk, 100 µM.)

 
Thrombin mediates intracellular signaling by binding with G protein-coupled PARs (7). To investigate the role of thrombin receptors in the thrombin-induced barrier protection in A549 cells, we utilized TRAP-6, a synthetic thrombin receptor–activating peptide composed of 6 amino acids (SFLLRN), which act as a PAR-1 agonist (7). The addition of SFLLRN caused a dose-dependent increase in TER across A549 cells (Figure 1D), as we have observed in thrombin-stimulated A549 cells. These results suggest that the barrier-protective effect of thrombin in A549 cells is mediated by PAR ligation.

Due to the critical role of the actin cytoskeleton in barrier regulation (10), we next examined the effect of thrombin on actin cytoskeleton reorganization using immunofluorescence microscopy. Consistent with our previous reports (11), thrombin (50 nM) induced the prominent formation of central stress fibers and intercellular gaps in the EC monolayers (Figure 2A). However, the actin cytoskeletal reorganization after thrombin stimulation of lung epithelial cells revealed enhanced cortical actin rearrangement abundant in the midplane of the cell monolayers (Figure 2A). Thus, our data suggest that thrombin induced a distinct pattern of actin rearrangement in the two human lung cell types, with the cortical actin remodeling in lung epithelium reminiscent of the actin rearrangement that we have observed in other barrier-protective stimuli, such as shear stress (15), hepatocyte growth factor (HGF) (17), and sphingosine 1-phosphate (S1P) (14).




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  		Figure 2. Actin cytoskeleton rearrangement and diphosphorylated MLC immunoreactivity in human pulmonary endothelial and epithelial cell monolayers. HPAEC and A549 monolayers were challenged with thrombin (50 nM) for 5 min. F-actin and ppMLC were detected by staining with Texas Red–conjugated phalloidin and anti-ppMLC–specific antibodies, which recognize phosphorylated serine19 and threonine18, respectively. (A) Actin staining in both cell types. In contrast with the predominant central stress fiber formation (arrowhead) and gap formation (arrows) in EC monolayers (upper panels), thrombin-challenged A549 monolayers revealed enhanced peripheral actin (arrowhead) in the basal plane (middle panels) and midplane (lower panels), with preservation of cytoskeletal architecture and monolayer integrity. (B) ppMLC immunoreactivity (arrowheads) correlates with the central stress fibers in thrombin-stimulated HPAEC, whereas MLC phosphorylation accumulates in the peripheral actin rim in A549 cells. Insets: cell monolayers at lower magnification.

 
Effects of Thrombin on MLC Phosphorylation: Role of MLCK- and RhoK-Dependent Barrier-Regulatory Mechanisms
Phosphorylation of MLC is a key determinant of EC contraction in several models of agonist-induced barrier dysfunction (10). This process is induced by Ca2+/calmodulin-dependent MLCK, and coordinates activation of the Rho/RhoK pathway and subsequent inhibition of MLC phosphatase. Total MLC levels in HPAEC and A549 total lysates were assessed by Western blotting, and our results suggest comparable amounts of total MLC expressed by these two cell types (data not shown). Thrombin-induced MLC phosphorylation in pulmonary epithelial and EC achieves similar peak MLC phosphorylation levels at ~ 5 min (Figure 3A). Thrombin-activated MLC phosphorylation in A549 was less robust, with ~ 1/3 of the magnitude in MLC phosphorylation compared with human endothelium. Interestingly, however, the total increase in MLC phosphorylation (fold increase from baseline) was similar between A549 and human EC (Figure 3A). The MLC phosphorylation in A549 cells declined rapidly to basal levels by 30 min, whereas ppMLC levels in HPAEC remained elevated at 30 min. Thrombin-induced MLC phosphorylation in both HPAEC and A549 was attenuated by preincubation, with either MLCK inhibitor (ML-7, 10 µM, 1 h) or RhoK inhibitor (Y-27632, 10 µM, 1 h), as detected by Western blot (Figure 3B). Immunofluorescence staining of ppMLC revealed distinct spatial distribution of ppMLC in the human endothelium and epithelium after thrombin exposure (Figure 2B). Consistent with actin reorganization, accumulation of ppMLC in A549 after thrombin challenge (5 min) appeared in the peripheral ring, whereas, in EC, ppMLC colocalized with the actin central stress fibers (Figure 2B). Finally, we assessed the effects of inhibition of MLC phosphorylation on thrombin-induced permeability of the HPAEC and A549 monolayer. Reductions in the levels of MLC phosphorylation, either by MLCK inhibitor (ML7) or RhoK inhibitor (Y-27632), attenuated thrombin response in both HPAEC and A549 cell lines (Figure 4). These results suggest divergent but critical roles of MLC phosphorylation in barrier-disruptive responses in EC and barrier-protective responses in epithelial cells.



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  		Figure 3. Role of MLCK and RhoK on thrombin-induced MLC phosphorylation in A549 and HPAEC. (A) The time course of MLC diphosphorylation to thrombin stimulation in HPAEC and A549 cultures was monitored by immunoblotting followed by calculations of the ratio of ppMLC/pan-MLC. Both HPAEC and A549 demonstrate a similar time course of thrombin-induced MLC phosphorylation, peaking at 5 min. Results of five independent experiments are expressed as mean ± SE (*P < 0.05, compared with vehicle controls). (B) Effect of MLCK and RhoK inhibitors on thrombin-induced MLC phosphorylation in HPAEC and A549. After 1-h preincubation with either MLCK inhibitor (ML-7, 10 µM) or RhoK inhibitor (Y-27632, 10 µM) followed by thrombin stimulation (50 nM, 5 min), MLC phosphorylation in HPAEC and A549 was assessed as described in MATERIALS AND METHODS. Effects of pharmacological inhibitors on MLC phosphorylation are expressed as percent of maximal phosphorylation induced by thrombin alone. Results of five independent experiments are expressed as mean ± SE (*P < 0.05 compared with thrombin alone). (Open bars, Thr; lightly shaded bars, Thr + ML-7; black bars, Thr + Y-27632.)

 


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  		Figure 4. Effects of MLCK and RhoK inhibitors on thrombin-induced TER changes. HPAEC (A and B) and A549 cells (C and D) were preincubated with either ML-7 (10 µM, 1 h), or Y-27632 (10 µM, 1 h) before thrombin stimulation. TER across EC and A549 monolayers was measured as described in MATERIALS AND METHODS. The concentrations of ML-7 (10 µM) and Y-27632 (10 µM) utilized to inhibit MLC phosphorylation in HPAEC have been described previously (14). Effective inhibitory concentrations (50 µM ML-7 and 50 µM Y-27632) for A549 cultures have been determined in preliminary experiments. Shown are results of five independent experiments. SE was < 0.03 ohms at each time point. Both ML-7 and Y-27632 significantly attenuated thrombin permeability responses in EC and A549 (P < 0.05). (A: diamond, vehicle; triangle, thrombin; square, ML-7 + Thr. B: diamond, vehicle; triangle, thrombin; square, Y-27632 + Thr. C: diamond, vehicle; square, thrombin; triangle, ML-7 + Thr. D: diamond, vehicle; square, thrombin; triangle, Y-27632 + Thr.)

 
Translocation of Tight Junction Proteins (ZO-1 and Occludin) and Adherens Junction Protein (ß-Catenin) after Thrombin Stimulation
Agonist-induced alterations in barrier function depend on the balance between contractile forces imposed by actomyosin stress fibers and the tethering forces imposed by peripheral actin ring and stabilized cell contact protein complexes, including tight junctions, adherens junctions, and focal adhesions (10). Figure 5A illustrates that EC express lower contents of the tight junction protein ZO-1 and occludin compared with epithelial cells (55 and 13%, respectively; P < 0.05), whereas ß-catenin levels are slightly increased in human endothelium (Figure 5A). We next analyzed the effects of thrombin on the stabilization of the tight-junction proteins, ZO-1 and occludin, and the adherens junction protein, ß-catenin. Cytosolic and membrane/cytoskeletal fractions were separated as described in MATERIALS AND METHODS, and the subcellular localization of occludin and ZO-1 in response to thrombin challenge was analyzed by immunoblotting. After thrombin challenge, we observed an ~ 2-fold decrease in ZO-1 content in the cytosolic fraction (29.3 ± 10% to 14.7 ± 8% of total protein content; P < 0.05) with parallel increase in membrane/cytoskeletal fraction (Figure 5B). Immunofluorescent staining of ZO-1 (Figure 6) suggests that the increased content of ZO-1 in the membrane/cytoskeletal fraction reflects ZO-1 translocation from cytoplasm to the functional cell contact sites, which potentially operate cytoskeletal/cytosolic linkage of cell contact sites in A549 after thrombin challenge. Biochemical and immunofluorescent assessment of occludin and ß-catenin in cytoskeletal/membrane fractions were not significantly altered in response to thrombin in either HPAEC or A549 (Figures 5B and 6). Finally, ZO-1, occludin, ß-catenin, and F-actin in thrombin-stimulated A549 appeared as a circumferentially-distributed ring in the middle plane of the cell monolayer (Figure 6). In contrast, EC junction proteins responded to thrombin challenge by disruption of the continuous peripheral pattern of ZO-1, occludin, and ß-catenin staining in association with increased intercellular gap formation.




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  		Figure 5. Differential expression and subcellular localization of ZO-1, occludin, and ß-catenin in HPAEC and A549 cells after thrombin. (A) The contents of ZO-1, occludin, and ß-catenin in total HPAEC and A549 cell lysates were detected by immunoblotting and quantified as described in MATERIALS AND METHODS. Tubulin was used for normalization of relative protein expression in HPAEC and A549. Results of five independent experiments are expressed as mean ± SE (*P < 0.05). Open bars, A549; black bars, HPAEC. (B) Differential effects of thrombin on subcellular localization of ZO-1, occludin, and ß-catenin in HPAEC and A549. Cytosolic and membrane/cytoskeletal protein fractions of thrombin-stimulated HPAEC and A549 were prepared as described in MATERIALS AND METHODS, and analyzed by immunoblotting. Tubulin was used as an internal control for sample loading. All blots were performed with the equal protein amounts in the samples. Western blots were quantified by scanning densitometry. Error bars represent ± 1 SE. Results of five independent experiments are expressed as mean ± SE (*P < 0.05).

 


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  		Figure 6. Effects of thrombin on intracellular localization of ZO-1, occludin, and ß-catenin in HPAEC and A549. HPAEC (A, B, E, F, I, J) and A549 (C, D, G, H, K, L) were challenged with thrombin (50 nM, 30 min) followed by immunofluorescent detection of ZO-1 (B and D), occludin (F and H), and ß-catenin (J and L). Arrows represent intercellular gaps in the HPAEC monolayer induced by thrombin stimulation. Arrowheads show discontinuous staining of ZO-1, occludin, and ß-catenin in HPAEC after thrombin challenge, whereas, in A549, ZO-1, occludin, and ß-catenin peripheral patterns were preserved. Control experiments for immunostaining without primary antibody were undertaken and revealed no fluorescent staining (data not shown).

 
Effect of Thrombin on Rho and Rac Activation in HPAEC and A549
The members of the Rho family of small GTP-binding proteins are key regulatory molecules associated with cytoskeleton changes and cell motility (18). Taking into account a significant role of small GTPases in cytoskeletal remodeling, we examined the activation of Rho and Rac in response to thrombin in these two cell types. Thrombin increased prominent Rho activity in both A549 and HPAEC, with peak activity at 5 min (Figures 7A and 7B). In contrast, Rac activation in response to thrombin was only observed in A549 (5 min), but not in HPAEC (Figures 7A and 7B), suggesting that the cell-specific barrier-regulatory response in A549 cells may involve both Rho- and Rac-mediated signaling pathways. We next utilized adenoviral infection of N17Rac1 to inhibit Rac 1 activity, and examined thrombin-induced barrier-protective responses in A549 cells. A dramatic increase in Rac expression, which reflects overexpression of DN Rac mutant, was observed in adenoviral-infected Rac DN cells (Figure 7C). The baseline TER was comparable in adenoviral-infected (control and N17Rac1) and noninfected A549 cells (data not shown). After thrombin challenge, we observed increased TER in A549 cells infected with control adenoviral vector containing green fluorescent protein and in noninfected A549 cells. In contrast, the barrier-protective response was significantly attenuated in N17Rac1-expressing A549 cells (Figure 7C). These data suggest a critical role for Rac-mediated molecular mechanisms in thrombin-induced barrier regulation in alveolar epithelial cells.



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  		Figure 7. Effect of thrombin on Rho and Rac activation in human pulmonary EC and epithelial cells. (A) HPAEC and A549 stimulated with thrombin (50 nM) for 0, 5, and 30 min. Rho-GTPase and Rac-GTPase activity was detected using pull-down Rho- and Rac-activation assays as described in MATERIALS AND METHODS and demonstrated by immunoblotting. GTP{gamma}S (10 mM) and GDP (100 mM) were used as positive (+) and negative (–) controls, respectively, for both Rho- and Rac-activation assay. (B) Results of quantitative analysis of Rac and Rho activation. Maximal thrombin-induced activation of Rho and Rac in A549 cells was taken as 100%. Rac- and Rho-GTPases are expressed in both cell types at comparable levels. (C) The effect of Rac molecular inhibition on the thrombin-induced TER increases in the A549 monolayer. The blot represents total Rac expression in noninfected cells, adenoviral-infected A549–expressing green fluorescent protein(AdV-GFP) as negative control, and adenoviral-infected A549 expressing DN Rac mutant (AdV-N17Rac). Adenoviral DN Rac mutant infection caused five-fold increases in Rac expression, as detected by Western blot, which was attributed to overexpression of DN Rac and downregulation of endogenous Rac activity. TER changes in response to thrombin stimulation (50 nM) were monitored as described in MATERIALS AND METHODS. After thrombin challenge, inhibition of Rac significantly blocked thrombin-induced barrier-protective response in A549 (P < 0.05). Shown are results of three independent experiments. SE was < 0.03 ohms at each time point. (Diamond, AdV-GFP + vehicle; triangle, AdV-GFP + Thr; square, AdV-N17Rac + Thr.)

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The lung alveolar epithelium forms a tight monolayer, which exists in intimate contact with lung capillary endothelium. Because both alveolar epithelial and capillary EC form a tight blood–gas barrier, preventing fluid leakage into the alveolar space, it is of primary importance to define the regulation of the endothelial and epithelial barrier integrity by bioactive edemagenic agonists. We have extensively analyzed the barrier-disruptive effects of thrombin on endothelial permeability and cytoskeletal rearrangements (6, 8, 11). In the present study, we examined thrombin effects on alveolar epithelial cell barrier regulation. In contrast to vascular endothelium, our results demonstrate that thrombin challenge produces alveolar epithelial cell barrier enhancement.

Thrombin-induced EC barrier disruption involves MLC phosphorylation and actomyosin contractile filament formation (11). We examined the potential roles of MLC phosphorylation and actin organization in A549 compared with HPAEC to explain the barrier-protective effect of thrombin on A549 monolayers. Strikingly divergent responses were observed, with thrombin inducing the formation of a peripheral actin ring containing phosphorylated MLC in pulmonary epithelial cells, whereas central stress fiber formation and loss of the thin cortical ring was observed in EC monolayers (Figure 2A and 2B). We have previously observed similar spatially-defined increases in actomyosin interaction, cortical actin thickening, and increased phosphorylated MLC in endothelium exposed to barrier-protective agents, such as HGF (17), S1P (14), and simvastatin (19), which correlated with increased TER and EC barrier enhancement. In the present study, we have extended these findings on the pulmonary alveolar epithelial cells and confirmed the requirement of Rac GTPase in actin ring formation and the maintenance of barrier integrity. The barrier-protective effect in A549 cells was attenuated by inhibition of MLCK and RhoK, suggesting that this response in A549 is MLCK- and Rho-dependent. The actin arrangement, MLCK activity, and MLC phosphorylation have been demonstrated as critical steps in gastrointestinal epithelial cell barrier regulation (20, 21). Importantly, we noted that the thrombin-induced barrier-protective response in epithelial cells is substantially prolonged compared with EC, an effect that is also observed with barrier-protective agents (14, 17, 19). Although speculation that the specific actin arrangement in each cell type is the important contributing factor that modifies the direction of contractile forces leading to different barrier regulation, our results suggest that MLC phosphorylation is not the sole factor responsible for thrombin-induced epithelial barrier regulation.

Cell junctions, in concert with the dynamic cytoskeleton, play an important role in cell–cell adhesion and barrier regulation (10, 13, 22). Specialized cell junctions form at multiple areas of cell–cell and cell–matrix contact in all tissues, but are particularly important and abundant in epithelial tissues (13). A functional unit, called apical junctional complex, is composed of the tight junctions and adherens junctions, which, together, structurally anastomose with the perijunctional actomyosin ring cytoskeleton and influence the maintenance of the permeability barrier that regulates paracellular transport in epithelial cells (23). We evaluated the role of tight-junction and adherens-junction proteins in response to thrombin in A549 and HPAEC, and found that one occludin-binding protein, ZO-1, translocates from the cytosolic compartment to the cell membrane contact sites after thrombin challenge in pulmonary epithelial A549 cells (Figures 5B and 6). The effects of thrombin on ZO-1 have not been documented in epithelial cells; however, ZO-1 interaction with the actin cytoskeleton either directly or via another actin-binding protein, cortactin (24), may be key to the barrier enhancement described in this study. Immunofluorescent staining demonstrated peripheral translocation of ZO-1 and its localization in close proximity to the enhanced actin ring (Figure 2A). These results may indicate specific actin remodeling in A549, which is, in contrast to HPAEC, driven by enhancement of peripheral tight junctions that anchor actin filaments and thus induce peripheral actin ring formation. The importance of ZO-1 in actin ring organization, which is generally accepted as a critical factor in determining epithelial paracellular permeability, has been recently demonstrated in human intestinal epithelial cells stimulated with interferon {gamma} (21). Using time-course analysis, Youakim and colleagues (21) showed that alterations in ZO-1 expression and apical actin perturbation induced by interferon-{gamma} could not be dissociated. The redistribution of ZO-1 to cell contact sites in the present study (Figures 5B and 6) may also maintain the stability of occludin and ß-catenin. The tight-junction transmembrane protein, occludin, is arranged in rows and forms contacts with neighboring cells, whereas cytosolic protein, ZO-1, interacts with the cytosolic domain of occludin (25) and creates a link with the cytoskeleton (13, 26). Importantly, the adhesiveness of occludin and, therefore, strength of tight junctions, correlates with occludin's ability to colocalize with its cytoplasmic binding protein, ZO-1. Furthermore, ZO-1 is believed to be responsible for coupling extracellular signaling pathways with the cytoskeleton. Our results support a critical role of ZO-1 in thrombin-mediated barrier regulation in alveolar epithelial cells; however, the correlation of ZO-1 and actin ring formation needs to be further elucidated.

The protein organization of the adherens junctions, which creates cell–cell adhesion just below the tight junctions, is analogous to the tight-junction proteins. Although the tight junctions and the adherens junctions work together as a single unit, interactions of tight-junction protein and adherens-junction protein are still unclear (27, 28). Because the localization of cadherin–catenin complex depends on the establishment of the cortical circumferential actin network (29), we speculate that MLCK-, Rho-, and Rac-mediated reorganization and peripheral F-actin ring enhancement in A549 cells promotes tight-junction and adherens-junction assembly and increases barrier integrity. Further studies are needed to fully delineate this interaction.

Thrombin interacts with PARs and triggers intracellular signaling pathways via activation of heterotrimeric G-proteins. Thrombin-mediated cleavage of the amino-terminal extension of PAR-1 unmasks a new amino terminal sequence, SFLLRN (TRAP6), which functions as a tethered peptide ligand and induces receptor activation and transmembrane signaling (7, 30). Our results demonstrate that both thrombin and TRAP6 induced significant TER increases in A549 cells, indicating PAR-1 involvement in thrombin-mediated barrier-protective response. The effects of thrombin on barrier integrity have been extensively demonstrated in association with Rho-GTPase activation (14, 31, 32). The Rho subfamily regulates stress fiber, tight-junction, and perijunctional actin formation (18). We have previously described the role of Rho in thrombin-induced stress fiber formation, MLC phosphorylation activation, and inhibition of MLC dephosphorylation associated with prolonged actomyosin contraction and endothelial barrier disruption (9, 32). In this study, we investigated the roles of the Rho family GTPases Rho and Rac in regulation of thrombin effects on pulmonary epithelial and EC, and demonstrated both Rho and Rac activation in A549 cells stimulated with thrombin. Moreover, both the inactivation of Rho by pharmacologic RhoK inhibitor (Y-27632), and Rac inhibition by adenoviral infection with DN Rac1 mutant blocked the barrier-protective effect of thrombin on A549 monolayers. Thus, our results support the role of both Rho and Rac activation in barrier enhancement of A549 cells.

Although Rho activation is essential for tight-junction assembly (13), the data regarding specific effects of Rho activation on the maintenance of the junctions are still controversial. Hopkins and colleagues (33) demonstrated that inhibition of RhoA activity by recombinant Clostridium botulinum toxin C3 induced disruption of perijunctional actin cytoskeleton, in concert with redistribution of ZO-1 and occludin away from tight-junction membranes and impaired tight-junction barrier function. Jou and coworkers (34) examined roles of RhoA and Rac1 in regulation of tight junctions by using GTPase mutants, and confirmed that inhibition of RhoA can disturb tight junction structure and function. Activation of Rho, Rac, and Cdc42 by Escherichia coli cytotoxic necrotizing factor-1 in T84 intestinal epithelial monolayers demonstrated reductions in TER, whereas cell pretreatment with RhoA inhibitor (C3 toxin) or RhoK inhibitor did not block the effects of cytotoxic necrotizing factor-1 (33). These results suggest that RhoA is not involved in reductions in TER or barrier function in T84 cells, and inhibition of RhoA or activation of Rac and Cdc42 may adversely affect that function. Other reports suggest that Rho activation plays a role in barrier disruption. Thus, the equilibrium of Rac, Rho, and Cdc42 Rho-GTPases activities may be essential for regulation of optimal barrier function in epithelial cells (33). Further studies will address precise mechanisms of this process.

Rac activation results in assembly of actin filaments at the cell periphery to produce lamellipodia and membrane ruffles, and induces accumulation of E-cadherin and ß-catenin at the cell–cell adhesion sites. We have previously demonstrated a strong association between Rac-GTPase and formation of cortical actin ring with barrier protective effects in EC after challenging with S1P (14), HGF (17), and simvastatin (19). In this study, thrombin-induced Rac-GTPase activation in A549 cells was observed early (5 min) and resulted in the formation of cortical actin ring. The mechanisms by which Rho-GTPases become activated through PAR-1 receptor in epithelial cell are not clear. Although thrombin activates Rho in the majority of cell types, including EC, it may also activate Rac and Cdc42 in human blood platelets (35, 36). Typically, in other cells, G-protein–coupled receptor-mediated activation of Rac has been shown to involve Gi-type G proteins. In EC, the G protein ß{gamma} subunits are involved in this process. Receptors, triggered by thrombin, couple to Gi, Gq, and G12/G13, and are able to rapidly recruit and activate several effectors, such as phospholipase C, protein kinase C, or phosphoinositide 3-kinase. We speculate that coupling of PAR-1 receptor with specific G proteins and downstream signal pathways in A549 has a pattern distinct from pulmonary EC, and may lead to Rac-GTPase activation in alveolar epithelial cells in response to thrombin stimulation. Therefore, the Rac and Rho interplay detected in thrombin-stimulated A549 may govern unique cytoskeletal rearrangements.

In summary, these results indicate, for the first time, the enhancement of barrier integrity by thrombin in alveolar epithelial cells, and describe the roles of actomyosin interaction, cell junction protein stability, and regulatory Rho-GTPases (Rho and Rac) in this response. Barrier protection, induced by thrombin in this immortalized neoplastic alveolar cell line, involving activation of Rho and Rac leads to MLC phosphorylation and formation of the peripheral actomyosin ring with peripheral accumulation of ZO-1/occludin complexes detected in A549 cells. We speculate that strong interaction of the peripheral actomyosin ring with enhanced tight-junction complexes may result in barrier enhancement observed in thrombin-stimulated A549 monolayers with ZO-1 (a key protein involved in the regulation of barrier function). If the enhancement of the alveolar epithelial barrier in response to thrombin can be extended to primary human alveolar epithelium, this paradoxical effect of thrombin may represent a compensatory mechanism that promotes the restoration of barrier integrity after acute lung injury.


   Acknowledgments
 
This work was supported by grants from the National Heart, Lung, and Blood Institute (HL 58064), Specialized Centers of Clinically Oriented Research in Acute Lung Injury (HL 73994), the Dr. David Marine Endowment, and a Faculty of Medicine award from Chulalongkorn University. The authors gratefully acknowledge the contributions of Lakshmi Natarajan for superb technical assistance and Denise Guise for expert administrative assistance with manuscript preparation.

Received in original form December 2, 2003

Received in final form July 2, 2004


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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2003-0432OCv1
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Proc. Am. Thorac. Soc. Am. J. Respir. Crit. Care Med.
Copyright © 2004 American Thoracic Society.