Tumor necrosis factor (TNF)-α, a major proinflammatory cytokine, triggers endothelial cell activation and barrier dysfunction which are implicated in the pathogenesis of pulmonary edema associated with acute lung injury syndromes. The mechanisms of TNF-α–induced vascular permeability are not completely understood. Our initial experiments demonstrated that TNF-α–induced decreases in transendothelial electrical resistance across human pulmonary artery endothelial cells are independent of myosin light chain phosphorylation catalyzed by either myosin light chain kinase or Rho kinase. We next assessed the involvement of another cytoskeletal component, the tubulin-based microtubule network, and found TNF-α to induce a decrease in stable tubulin content and partial dissolution of peripheral microtubule network as evidenced by anti-acetylated tubulin and anti–β-tubulin immunofluorescent staining, respectively. Microtubule-stabilizing agents, paclitaxel and epothilone B, significantly attenuated TNF-α–induced decreases in transendothelial electrical resistance, inhibited the cytokine-induced increases in actin stress fibers, formation of intercellular gap, and restored the TNF-α–compromised vascular endothelial (VE)-cadherin–based cell–cell junctions. Importantly, neither TNF-α nor paclitaxel treatment was associated with endothelial cell apoptosis. Inhibition of p38 mitogen-activated protein kinase by SB203580 significantly attenuated TNF-α–induced microtubule destabilization, actin rearrangement, and endothelial barrier dysfunction. These results strongly suggest the involvement of microtubule rearrangement in TNF-α–induced endothelial cell permeability via p38 mitogen-activated protein kinase activation.
Disruption of the vascular barrier is a prominent feature of acute lung injury syndromes, and results in pulmonary edema formation and subsequent respiratory dysfunction (1). Endothelial cell activation is often accompanied by enhanced cellular contraction and formation of intercellular gaps, major events leading to pulmonary interstitial edema (2). The endothelial barrier integrity is the result of a balance between the tethering and contractile forces acting on endothelial cells, which are critically dependent upon cytoskeletal components, including the actin-based microfilaments, intermediate filaments, and microtubules (3). Tumor necrosis factor (TNF)-α, a proinflammatory cytokine secreted by macrophages and endothelial cells, has been implicated in endothelial cell activation, increased endothelial cell permeability, and pulmonary edema formation (4, 5). However, the exact mechanisms by which TNF-α triggers vascular barrier dysfunction are not precisely defined. We and others have noted that TNF-α induces endothelial actin microfilament rearrangement and intercellular gap formation that parallel the development of transendothelial permeability (5, 6). It is less clear which signaling pathways are orchestrating these morphologic changes of endothelial cells and their relative direct contribution to the vascular barrier dysfunction. We have previously shown that TNF-α induces significant actin microfilament rearrangement by myosin light chain phosphorylation, as a consequence of the coordinated action of the myosin light chain kinase (MLCK) and Rho kinase (5). However, TNF-α–induced endothelial cell barrier dysfunction was shown to evolve in an MLCK- and Rho kinase–independent fashion (5), suggesting that MLCK-independent microfilament changes and/or other cytoskeletal structures, such as intermediate filaments, microtubules, and adherens junctions may be involved. Although it was previously shown that TNF-α triggers microtubule disassembly (7), the role of the microtubule cytoskeleton in TNF-α–induced pulmonary endothelial permeability is largely unknown. There is, however, encouraging evidence that the microtubule network may be an important modulator of endothelial barrier function using other agonists, such as nocodazole (8), light-exposure (9), and hydrogen peroxide (10). In the present report, we investigated the role of MLCK-independent regulation of TNF-α–induced endothelial cell cytoskeletal changes and permeability. Our results indicate that microtubule destabilization contributes to both TNF-α–induced endothelial cell actin microfilament cytoskeletal changes and barrier dysfunction via a p38–mitogen-activated protein kinase–(MAPK)-dependent pathway.
Human pulmonary artery endothelial cells, purchased from Clonetics (BioWhittaker, Inc., Walkersville, MD) were used for experiments at passages 7–9. The cells were maintained in complete culture medium consisting of 20% bovine serum, endothelial cell growth supplement (17 μg/ml, H-neurext; Upstate Biotechnology, Lake Placid, NY), and penicillin/ streptomycin (100 U/ml; Gibco, Invitrogen Corporation, Carlsbad, CA) at 37oC in an atmosphere of 5% CO2 and 95% air. TNF-α (biological activity of 2 × 107 U/mg) and paclitaxel were purchased from Sigma Aldrich (St. Louis, MO). Texas Red-X phalloidin was purchased from Molecular Probes, Inc. (Eugene, OR). SB203580 and epothilone B were from Calbiochem-Novalbiochem Corp. (La Jolla, CA). The anti-VE cadherin antibody was from BD Biosciences–Transduction Laboratories (Lexington, KY), the anti-acetylated tubulin antibody was from Accurate Chemicals (Westbury, NY), the anti–phospho-p38 MAPK and anti-p38 MAPK antibodies were from Cell Signaling Technology (Beverly, MA), and anti–β-tubulin antibody was from ICN Diagnostics (Orangeburg, NY).
Human pulmonary artery endothelial cells were cultured to confluence in 12-well dishes on coverslips coated with gelatin. After exposure to agonists, endothelial cell monolayers were fixed in 3.7% formaldehyde and then permeabilized with 0.25% Triton X-100. After staining, coverslips were mounted on slides and examined under oil-immersion (60× or 100× magnification) using an Eclipse TE300 inverted microscope (Nikon Inc., Melville, NY). Actin was visualized by Texas Red-phalloidin staining (1:200) for 1 h at room temperature. This method enabled the examination of endothelial cell morphology, intercellular gap formation in confluent monolayers, and intracellular actin filament reorganization (stress fiber formation, cortical or perinuclear actin organization). Staining for VE-cadherin, acetylated tubulin, or β-tubulin was performed in a similar manner, with 1 h incubation at room temperature with the primary antibody, followed by three washes with PBS-Tween (0.1%) and incubation for 1 h at room temperature with an appropriate secondary antibody conjugated to immunofluorescent dye (Alexa 488; Molecular Probes, Inc., Eugene, OR). After three washes with PBS-Tween, the coverslips were mounted and analyzed using Nikon video-imaging system (Nikon, Inc.) consisting of phase contrast inverted microscope connected to digital camera attached to image processor and images were recorded and saved in an Adobe Photoshop 4 program (Adobe Systems, Inc., San Jose, CA), using Pentium II PC.
Endothelial cell proteins were separated by SDS-PAGE, transferred to Immobilon PVDF membrane (Millipore, Bedford, MA), and immunoblotted for 1 h with the primary antibody as previously described (5), followed by the addition of the appropriate horseradish peroxidase–conjugated secondary antibody (1:10,000). The reaction was visualized by enhanced chemiluminescence (ECL) and autoradiography (Amersham Biosciences, Inc., Piscataway, NJ), according to the manufacturer's instructions.
Quantitation of apoptotic endothelial cells was obtained using Nucleosome ELISA Kit (Oncogene Research Products, Cambridge, MA), following the manufacturer's protocol. In these experiments, endothelial cells were lysed and the supernatant loaded onto precoated DNA-binding protein wells. The nucleosomes were detected using anti-histone 3 biotinylated antibody followed by streptavidin horseradish peroxidase with absorbance (450 nm) compared to lyophilized standards with designated nucleosome unit values. The data are expressed as a mean index relative to the vehicle-treated control. Two experiments were performed and for each independent experiment the samples were loaded in duplicate.
Electrical resistance of human pulmonary artery endothelial cell monolayer was measured using electrical cell impedance sensor technique as we have previously described (5). In this system (Applied Biophysics, Inc., Troy, NY), endothelial cells are cultured on a small gold electrode (10-4 cm2) in complete media. The endothelial monolayer acts as insulating particles and the total resistance across the monolayers is composed of the resistance between the ventral cell surface and the electrode and the resistance between cells. A 4,000-Hz AC signal with 1 V amplitude through a 1 mΩ resistor created an approximate constant current source (1 μA). The lock-in amplifier attached to the electrodes detects changes in both magnitude and phase of the voltage appearing across the endothelial cell and was controlled by an IBM-compatible personal computer, which was used both for data accumulation and processing. TER increased immediately after cell attachment and achieved a steady state when endothelial cells became confluent. Thus, experiments were conducted after the electrical resistance achieved a steady state. Resistance data was normalized to the initial voltage and plotted as a normalized TER. Only wells in which the TER achieved > 5,000 ohms were utilized.
In endothelial cells, the microtubules organize into a fine lattice network that spans the entire cytoplasm extending to the cell periphery (Figure 1Aa, c) (8). Treatment with TNF-α induces disassembly of peripheral microtubule network, as assessed by β-tubulin and acetylated tubulin immunostaining (Figure 1Ab, d). TNF-α–induced changes in microtubule structure, consisting of a loss of peripheral staining for acetylated (stable) tubulin (11), are detected as early as 3 h by microscopy (Figure 1Ab), and 1 h by Western blotting (Figure 1B). These changes precede significant TNF-α–induced increases in endothelial cell permeability seen typically at 4-5 h, as measured by TER (Figure 2A) , or albumin clearance across endothelial monolayers grown on filters (5, 6). β-Tubulin staining demonstrates that TNF-α–induced disassembly of total tubulin network (maximal at 16 h, Figure 1Ad) coincides with maximal TNF-α–induced increase in permeability, measured at 10 h by TER (Figure 2A).
We examined TNF-α–induced changes in endothelial cell permeability utilizing the electrical cell impedance sensor technique, a method extensively validated as a surrogate measurement of permeability for the endothelial monolayer (12–14), in which endothelial cells are grown to confluence on gold microelectrodes and TER is measured and expressed as normalized resistance to the initial measured voltage. TNF-α caused significant time- and dose-dependent decreases in TER beginning at 4–5 h of exposure, reaching a maximum decline at 10 h with changes persisting for 24–48 h (Figure 2A) (5). Pretreatment of endothelial cells with the known microtubule-stabilizing agent paclitaxel led to a significant attenuation (45.4% ± 10) in TNF-α–induced TER decrease, suggesting a role for microtubule destabilization in TNF-α–induced endothelial cell permeability. The onset of paclitaxel's protective effects on TNF-α–perturbed TER occurred within 3 h of cytokine treatment (range 2–5 h), consistent with an effect on microtubule disorganization which preceded the TER changes. This result was confirmed with epothilone B (Figure 2B), a synthetic analog of naturally occurring epothilones extracted from a strain of Myxobacterium sporangium which induces potent tubulin assembly and stabilization, similar to paclitaxel (15). Epothilone B induced similar magnitudes of TER attenuation of the maximal TNF-α response (39.2% ± 6.8, Figure 2C) when compared with paclitaxel. However, the precise mechanisms by which epothilone B stabilizes microtubules may differ form paclitaxel, suggested by differences seen in the shape of the effect on TNF-induced TER (Figures 2A and 2B). The attenuation in TNF-α–induced permeability was dose-dependent for both agents, with maximal effects obtained with 10 μM of paclitaxel, and 0.1 μM epothilone B, respectively (data not shown). Because both TNF-α and microtubule-stabilizing agents are recognized inducers of apoptosis and may act synergistically in various tumors (16), we evaluated whether they are inducing apoptosis in human pulmonary artery endothelial cells. Neither TNF-α nor paclitaxel, at the concentrations and in the conditions (20% serum) evaluated in our endothelial permeability model, are associated with significant induction of programmed cell death (Figure 2D), suggesting a limited role of apoptosis in TNF-α–induced microtubule remodeling and increase in permeability.
The integrity of the endothelial cell barrier function is the result of a balance between contractile and tethering forces that act on the endothelium. We have previously demonstrated that the TNF-α–triggered decreases in pulmonary TER are temporally associated with profound actin cytoskeletal rearrangement and cellular contraction (5). In endothelial cell from other vascular beds (human umbilical veins or mesenteric veins), it has been also shown that TNF-α induces disruption in VE-cadherin cellular distribution, thus decreasing tethering forces on the monolayer (17, 18). We next investigated whether there is a crosstalk between the microtubule and the actin cytoskeleton. Treatment of endothelial cell with paclitaxel alone (Figure 3)did not induce significant changes compared with control (not shown), the cells maintaining their polygonal shape, with predominant actin cortical staining and tight intercellular contacts with uniform VE-cadherin distribution. TNF-α (20 ng/ml, 3 h) induces actin stress fiber formation, cellular contraction, intercellular gap formation, and disruption of cell–cell contact with loss of VE-cadherin staining at the cell periphery (Figure 3). Microtubule stabilization with paclitaxel before TNF-α treatment resulted in a dramatic inhibition of the TNF-α−induced changes on the cytoskeleton, with marked reductions in the amount of stress fibers and preservation of the cellular shape, intercellular contacts, and VE-cadherin distribution and staining pattern (Figure 3). These results suggest than TNF-α–triggered microtubule disassembly is critical for most of the actin cytoskeletal and VE-cadherin changes.
Exposure of human pulmonary artery endothelial cells to TNF-α (20 ng/ml) resulted in an early (5 min) and robust increase in p38 MAPK phosphorylation consistent with enzymatic activation (Figure 4A). Both the time-course of the TNF-α–induced p38 MAPK activation and the lack of inhibition in the paclitaxel-pretreated cells (Figure 4B) indicate that TNF-α–induced p38 activation precedes the microtubule disassembly. To explore the role of p38 MAPK in endothelial cell permeability and actin and microtubule cytoskeletal rearrangement, we used a pharmacologic approach with a specific p38 MAPK inhibitor, SB203580 (20 μM, 45 min). As seen in Figure 4A, this inhibition completely abolished TNF-α–induced p38 MAPK activation. Pretreatment with SB203580 consistently resulted in significant inhibition of TNF-α–induced permeability (Figure 5A) with an average of 68% inhibition (Figure 5B), whereas SB203580 alone fails to cause any significant change over control in TER (Figure 5A). Consistent with the effects of SB203580 on TNF-α–induced barrier dysfunction, inhibition of p38 MAPK also prevented the TNF-α–induced microtubule disassembly (detected with acetylated tubulin staining) and endothelial cell actin rearrangement (Figure 6) . Pretreatment with SB203580 alone had no significant impact on basal microtubule and actin cytoskeletal staining (not shown). These results suggest a novel critical role of p38 MAPK activation in TNF-α–induced microtubule remodeling, actin cytoskeletal changes, and subsequent endothelial cell barrier dysfunction, as shown in Figure 7 .
TNF-α is a proinflammatory cytokine produced by activated leukocytes and endothelial cells resulting in upregulation of endothelial adhesion molecules, alterations in endothelial cell permeability (4, 5), and increased edema in isolated perfused lungs (19). The endothelial cell barrier function is maintained by the balance between the tethering and contractile forces acting on endothelial cells, which are critically dependent upon cytoskeletal components, including the actin-based microfilaments, intermediate filaments, and microtubules. TNF-α–induced endothelial cell activation includes actin cytoskeletal rearrangement, with an increase in actin stress fiber formation followed by intercellular gaps formation (5, 17, 20, 21). We have previously demonstrated that although TNF-α triggers MLCK and Rho kinase–dependent actin cytoskeletal rearrangements, the endothelial cell permeability is unaltered by inhibitors of these two enzymes.
Thus, the mechanism of TNF-α–triggered endothelial barrier dysfunction may involve MLCK-independent microfilament changes and/or other cytoskeletal structures, such as intermediate filaments, microtubules, and adherens junction proteins. In this study we evaluated the role of the endothelial cell microtubule network in TNF-α–induced barrier dysfunction. We found that TNF-α destabilizes the endothelial cell microtubule network concomitant with the onset of increased endothelial permeability, and that prevention of microtubule assembly significantly inhibits the effect of TNF-α on the endothelial barrier function. We next showed that microtubule stabilization inhibited actin rearrangement, VE-cadherin redistribution, and intercellular gap formation in response to TNF-α, suggesting important crosstalk between the microtubule cytoskeleton, the actin microfilaments, and the zonula adherens proteins. Although it has been previously reported that TNF-α leads to changes in the microtubule network (7), there is limited information available as to how the microtubule network may affect cellular shape and its contractile properties. Danowki et al found that microtubule disassembly led to an increase in nonmuscle (fibroblast) cell contractility and focal adhesion assembly, which was prevented by microtubule stabilization (22). Our laboratory has found that nocodazole-induced microtubule disruption leads to MLCK-independent, but Rho kinase–dependent MLC phosphorylation, actin rearrangement, and increased permeability in bovine pulmonary artery endothelial cells (8), suggesting that the microtubules are opposing cellular contraction.
In response to TNF-α, however, the mechanisms by which microtubule disassembly leads to permeability are likely to involve additional pathways, because the TNF-α–induced decreases in TER are independent of Rho and Rho kinase activation (5). One mechanism may involve the zonula adherens component VE-cadherin, because microtubule stabilization prevented the disruption in VE-cadherin staining, suggesting that TNF-α–induced microtubule disassembly triggers changes in VE-cadherin subcellular distribution, potentially weakening the intercellular junctions. Although microtubule disassembly may modulate TNF-α–induced cellular responses through additional mechanisms, these are less likely to be significant for our model. For example, an intact microtubule network was reported to be important for the maintenance of the steady-state of TNF-α receptor (23), hence its disruption leading to a downregulation of the TNF-α response, which is not observed in our permeability assays. Also, it is known that in several cell types microtubule stabilization (paclitaxel) leads to apoptosis, and this effect is synergistic to TNF-α–induced apoptosis (24). In our model, however, apoptosis is not a prominent feature (Figure 2D). We have also previously reported that TNF-α–induced permeability develops independently from apoptosis (5). The dichotomy between apoptosis and permeability is also suggested by similar magnitudes and kinetics of permeability in human and bovine pulmonary endothelial cells exposed to TNF-α, although their sensitivities to TNF-α–induced apoptosis are different. In standard culture conditions human pulmonary artery endothelial cells are resistant to TNF-α–induced apoptosis (Figure 2D) (25), whereas bovine pulmonary artery endothelial cells exhibit enhanced apoptosis in response to TNF-α (5, 26). TNF-α actually activates the pro-survival Akt pathway in human endothelial cells (27), explaining diminished apoptosis levels when compared with background control. A similar phenomenon of increase in pulmonary endothelial cell permeability independent of apoptosis has also been reported in response to lipopolysaccharide (28).
We showed that TNF-α triggers microtubule network disassembly, and that the microtubule cytoskeleton mediates TNF-α–induced pulmonary endothelial cell permeability through cellular shape changes involving the actin cytoskeleton and the zonula adherens proteins. The precise mechanisms by which TNF-α leads to this sequence of events are not well understood. The MAPK pathways are important intracellular signaling pathways that are activated by TNF-α–induced ligation of the TNF receptors TNFR1 (p55) and, to a lesser extent, TNFR2 (p75). Each component of the MAPK pathways is differentially recruited by specific stimuli resulting in kinase-specific signaling and regulation of cell growth, differentiation, and apoptosis. We found that p38 MAPK is a critical mediator of TNF-α–induced microtubule disassembly, actin microfilament rearrangement, and endothelial permeability. The p38 MAPK is a stress-activated MAPK family that consists of four isoforms: p38-α, p38-β, p38-γ, and p38-δ. p38-α and -β are inhibited by cytokine-suppressive anti-inflammatory drugs, the prototype of which is the compound SB203580, which we used in our studies. How p38 MAPK regulates microtubule dynamics in not well understood, but may involve direct phosphorylation of microtubule-associated proteins. For example, tau is a substrate for p38-γ, and its phosphorylation resulted in a marked reduction of its ability to promote microtubule assembly (29); and stathmin, a cytoplasmic protein linked to regulation of microtubule dynamics, is a substrate for p38-δ (30). In addition, p38 MAPK inactivates kinesin, an ATPase that mediates plus end–directed transport of organelles along microtubules (31). Although we do not know yet the precise mechanism by which p38 MAPK leads to microtubule remodeling in our model, our work suggests that this pathway is of critical importance in TNF-α–induced endothelial cell permeability. An important role for p38 MAPK in endothelial permeability has been previously implicated in TNF-α–induced permeability in human umbilical vein endothelial cells (32, 33) and also in response to VEGF (34) and hydrogen peroxide (35), but the precise mechanisms by which p38 MAPK leads to endothelial cell barrier dysfunction are not known. Given the more effective inhibition of TNF-α–induced barrier dysfunction with the SB203580 compared to the microtubule stabilizing agents, it is likely that the downstream effects of p38 MAPK activation encompass multiple cellular pathways, in addition to the microtubule cytoskeleton (schematic in Figure 7). The actin cytoskeleton function may be regulated by p38 MAPK via direct activation of MAPK-activated protein kinase-2 (MAPKAP kinase-2), which in turn phosphorylates and activates an actin-binding protein, heat shock protein 27 (HSP-27). Nonphosphorylated HSP-27 normally exists in high molecular weight multimers that serve as chaperones. Serine phosphorylation of HSP-27 results the dissociation of HSP-27 into monomers and dimers with redistribution of HSP-27 to the actin cytoskeleton and subsequent actin reorganization into stress fibers (36–39).
In conclusion, our results indicate that microtubule destabilization contributes to both TNF-α–induced endothelial cell actin microfilament cytoskeletal changes and barrier dysfunction. We also showed that p38-MAPK activation is a critical signaling event in the TNF-α–induced endothelial barrier dysfunction, by regulating the microtubule and actin cytoskeleton remodeling. Future work will help understand the precise mechanisms by which the MAPK signaling pathways regulate cytoskeletal functions. Elucidation of the complex orchestration of the endothelial barrier function will lead to potential therapies for the pulmonary edema seen in acute lung injury syndromes.
This work was supported by grants from the National Heart, Lung Blood Institute (HL 04396 [I.P.], HL 67307, HL 68062 [A.D.V.], HL 58064 [J.G.N.G.]), and a grant from the American Heart Association (A.D.V.). The authors gratefully acknowledge the contributions of Steve Durbin and Lakshmi Natarajan for superb technical assistance.
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