Abstract
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 1
Figure 1. Effect of TNF-α on the microtubule network in human pulmonary artery endothelial cells. (A) Photomicrographs of endothelial cells stained for stable tubulin with acetylated tubulin antibody (a and b) and for microtubules with β-tubulin antibody (c and d) and visualized with fluorescent microscopy. In control conditions (a and c, and insets), the endothelial cell microtubule network is composed of a fine network of stable microtubule structures. Treatment with TNF-α (20 ng/ml) results in microtubule disassembly, with early loss of peripheral stable tubulin (c and inset; 3 h) and loss of the elongated network structures (d and inset; 16 h). (B) Immunoblots of endothelial cells exposed to TNF-α (20 ng/ml; 1 h) or the microtubule stabilizing agent, paclitaxel (10 μM; 1 h) and blotted for acetylated tubulin or β-tubulin. Compared with vehicle-treated cells, TNF-α induced a decrease in stable tubulin, as identified by acetylated tubulin.
[More] [Minimize]
Figure 2. Effect of microtubule-stabilizing agents on TNF-α–induced endothelial permeability, as measured by TER and apoptosis. A and B show representative tracings (n = 4) of normalized electrical resistance measured across endothelial cell monolayers. Endothelial cells were exposed to vehicle (control) and TNF-α (20 ng/ml), where the gray arrow on the abscissa indicates the time of challenge. TNF-α–induced TER reduction was evident at 4 h, with a maximal effect at 10 h of exposure. (A) Microtubule stabilization by paclitaxel (10 μM; 1 h; black arrow) significantly inhibits TNF-α–induced decreases in resistance. Paclitaxel alone does not cause a sustained decrease in TER. (B) Microtubule stabilization by epothilone B (0.1 μM; 1 h; black arrow) also attenuates the TNF-α–induced decline in TER, whereas the drug alone has no effect on the baseline TER. (C) Bar graph demonstrating the effect of the microtubule stabilizing agents paclitaxel end epothilone B on the maximal TNF-α–induced decrease in TER. No significant difference was noted between the two agents (P = 0.44). (D) Bar graph indicating the apoptotic index calculated by nucleosomal ELISA. Human pulmonary artery endothelial cells maintained in standard culture conditions (20% serum) and exposed to TNF-α (20 ng/ml; 16 h), paclitaxel (10 μM; 16 h), or their combination exhibited no significant apoptosis. Bovine pulmonary artery endothelial cells exposed to TNF-α (20 ng/ml; 16 h) are shown as a positive control.
[More] [Minimize]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)
Figure 3. Outcome of microtubule stabilization on TNF-α–induced endothelial cell microfilament rearrangement and VE-cadherin subcellular distribution. Photomicrographs of endothelial cells exposed to paclitaxel (10 μM; 3.5 h), TNF-α (20 ng/ml; 3 h), or both and stained for actin with Texas Red phalloidin (upper panels, red) or for both actin and VE-cadherin (lower panels: actin, red; VE-cadherin, green) and visualized with fluorescent microscopy. Microtubule stabilization inhibits TNF-α–induced microfilament rearrangement that leads to cell shape changes, intercellular gap formation (arrowheads), and compromise in VE-cadherin intercellular contacts (arrows), with little effect on stress fiber formation.
[More] [Minimize]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)
Figure 4. The role of p38-MAPK activation in TNF-α–induced microtubule remodeling. Immunoblots of endothelial cells exposed to TNF-α (20 ng/ml) for the indicated times with or without a p38 MAPK inhibitor, SB203580 (A), or a microtubule stabilizing agent, paclitaxel (B), using both an anti–phospho-specific p38 MAPK antibody and an anti-p38 MAPK antibody. TNF-α–induced rapid p38 MAPK activation that was not attenuated by paclitaxel pretreatment, suggesting that p38 MAPK activation precedes the microtubule disassembly.
[More] [Minimize]
Figure 5. The effects of p38 MAPK inhibition on TNF-α–induced endothelial permeability. (A) Representative tracing (n = 4) of normalized TER measured across endothelial cell monolayers. Endothelial cells were exposed to vehicle (CTL) and TNF-α (20 ng/ml), where the gray arrow on the abscissa indicates the time of challenge. p38 MAPK inhibition by SB203580 (20 μM) pretreatment (black arrow indicates starting time) dramatically inhibits TNF-α–induced decreases in TER, indicating that p38 MAPK activation is critical for TNF-α–induced endothelial permeability. SB203580 alone does not cause changes in TER. (B) Bar graph indicating the % change in TER-induced by TNF-α in the presence of vehicle or SB203580 pretreatment. Bars represent standard deviation, and asterisk denotes statistical significance with a P = 0.01 by Student's t test.
[More] [Minimize]
Figure 6. The effect of p38 MAPK inhibition on the TNF-α–induced microtubule disassembly and actin cytoskeletal rearrangement. Photomicrographs of endothelial cells stained for actin (red) with Texas Red phalloidin and acetylated tubulin (green) and visualized with fluorescent microscopy. (A) Control conditions (complete culture medium, 4 h). Note the tight intercellular contact and the predominantly peripheral cortical actin staining (arrowhead). (B) TNF-α (20 ng/ml, 4 h) causes intercellular gap formation (arrows) and marked increases in actin stress fibers. (C) Pretreatment with the p38 MAPK inhibitor, SB203580, abolishes the changes seen with TNF-α, with less extensive stress fiber formation. (D) In control conditions the acetylated tubulin staining indicates intact microtubule structures forming a fine cytoplasmic lattice that extends to the cell periphery (inset). TNF-α triggers disassembly of these structures, with disappearance of acetylated tubulin staining at the cell periphery (E, inset). Pretreatment with the p38 inhibitor SB203580 prevents the disruptive effects of TNF-α on microtubules (F), with maintenance of the peripheral acetylated tubulin staining (inset), indicating that the TNF-α–induced microtubule and actin cytoskeletal changes are p38 MAPK-dependent.
[More] [Minimize]
Figure 7. Pathways of TNF-α–triggered endothelial cell permeability. TNF-α acts via p38 MAPK to trigger microtubule disassembly and subsequent actin microfilament rearrangement and cell-cell contact disruption. The interrupted lines reflect additional microtubule-independent cytoskeletal mechanisms of barrier dysfunction that may be regulated by p38 MAPK.
[More] [Minimize]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.
| 1. | Tomashefski, J. F., Jr. 2000. Pulmonary pathology of acute respiratory distress syndrome. Clin. Chest Med. 21:435–466. |
| 2. | Stevens, T., J. G. Garcia, D. M. Shasby, J. Bhattacharya, and A. B. Malik. 2000. Mechanisms regulating endothelial cell barrier function. Am. J. Physiol. Lung Cell. Mol. Physiol. 279:L419–L422. |
| 3. | Dudek, S. M., and J. G. Garcia. 2001. Cytoskeletal regulation of pulmonary vascular permeability. J. Appl. Physiol. 91:1487–1500. |
| 4. | Goldblum, S. E., B. Hennig, M. Jay, K. Yoneda, and C. J. McClain. 1989. Tumor necrosis factor alpha-induced pulmonary vascular endothelial injury. Infect. Immun. 57:1218–1226. |
| 5. | Petrache, I., A. D. Verin, M. T. Crow, A. Birukova, F. Liu, and J. G. Garcia. 2001. Differential effect of MLC kinase in TNF-alpha-induced endothelial cell apoptosis and barrier dysfunction. Am. J. Physiol. Lung Cell. Mol. Physiol. 280:L1168–L1178. |
| 6. | Goldblum, S. E., and W. L. Sun. 1990. Tumor necrosis factor-α augments pulmonary arterial transendothelial albumin flux in vitro. Am. J. Physiol. 258:L57–L67. |
| 7. | Molony, L., and L. Armstrong. 1991. Cytoskeletal reorganizations in human umbilical vein endothelial cells as a result of cytokine exposure. Exp. Cell Res. 196:40–48. |
| 8. | Verin, A. D., A. Birukova, P. Wang, F. Liu, P. Becker, K. Birukov, and J. G. Garcia. 2001. Microtubule disassembly increases endothelial cell barrier dysfunction: role of MLC phosphorylation. Am. J. Physiol. Lung Cell. Mol. Physiol. 281:L565–L574. |
| 9. | Sporn, L. A., and T. H. Foster. 1992. Photofrin and light induces microtubule depolymerization in cultured human endothelial cells. Cancer Res. 52:3443–3448. |
| 10. | Hinshaw, D. B., J. M. Burger, B. C. Armstrong, and P. A. Hyslop. 1989. Mechanism of endothelial cell shape change in oxidant injury. J. Surg. Res. 46:339–349. |
| 11. | Schulze, E., D. J. Asai, J. C. Bulinski, and M. Kirschner. 1987. Posttranslational modification and microtubule stability. J. Cell Biol. 105:2167–2177. |
| 12. | Furie, M. B., E. B. Cramer, B. L. Naprstek, and S. C. Silverstein. 1984. Cultured endothelial cell monolayers that restrict the transendothelial passage of macromolecules and electrical current. J. Cell Biol. 98:1033–1041. |
| 13. | Kazakoff, P. W., T. R. McGuire, E. B. Hoie, M. Cano, and P. L. Iversen. 1995. An in vitro model for endothelial permeability: assessment of monolayer integrity. In Vitro Cell. Dev. Biol. Anim. 31:846–852. |
| 14. | Garcia, J. G., K. L. Schaphorst, S. Shi, A. D. Verin, C. M. Hart, K. S. Callahan, and C. E. Patterson. 1997. Mechanisms of ionomycin-induced endothelial cell barrier dysfunction. Am. J. Physiol. 273:L172–L184. |
| 15. | Nicolaou, K. C., N. Winssinger, J. Pastor, S. Ninkovic, F. Sarabia, Y. He, D. Vourloumis, Z. Yang, T. Li, P. Giannakakou, and E. Hamel. 1997. Synthesis of epothilones A and B in solid and solution phase. Nature 387:268–272. |
| 16. | Nimmanapalli, R., C. L. Perkins, M. Orlando, E. O'Bryan, D. Nguyen, and K. N. Bhalla. 2001. Pretreatment with paclitaxel enhances apo-2 ligand/tumor necrosis factor-related apoptosis-inducing ligand-induced apoptosis of prostate cancer cells by inducing death receptors 4 and 5 protein levels. Cancer Res. 61:759–763. |
| 17. | Wojciak-Stothard, B., A. Entwistle, R. Garg, and A. J. Ridley. 1998. Regulation of TNF-alpha-induced reorganization of the actin cytoskeleton and cell-cell junctions by Rho, Rac, and Cdc42 in human endothelial cells. J. Cell. Physiol. 176:150–165. |
| 18. | Wong, R. K., A. L. Baldwin, and R. L. Heimark. 1999. Cadherin-5 redistribution at sites of TNF-alpha and IFN-gamma-induced permeability in mesenteric venules. Am. J. Physiol. 276:H736–H748. |
| 19. | Hocking, D. C., P. G. Phillips, T. J. Ferro, and A. Johnson. 1990. Mechanisms of pulmonary edema induced by tumor necrosis factor-alpha. Circ. Res. 67:68–77. |
| 20. | Brett, J., H. Gerlach, P. Nawroth, S. Steinberg, G. Godman, and D. Stern. 1989. Tumor necrosis factor/cachectin increases permeability of endothelial cell monolayers by a mechanism involving regulatory G proteins. J. Exp. Med. 169:1977–1991. |
| 21. | Goldblum, S. E., X. Ding, and J. Campbell-Washington. 1993. TNF-alpha induces endothelial cell F-actin depolymerization, new actin synthesis, and barrier dysfunction. Am. J. Physiol. 264:C894–C905. |
| 22. | Danowski, B. A. 1989. Fibroblast contractility and actin organization are stimulated by microtubule inhibitors. J. Cell Sci. 93:255–266. |
| 23. | Galeotti, T., D. Boscoboinik, and A. Azzi. 1993. Regulation of the TNF-alpha receptor in human osteosarcoma cells: role of microtubules and of protein kinase C. Arch. Biochem. Biophys. 300:287–292. |
| 24. | Lanni, J. S., S. W. Lowe, E. J. Licitra, J. O. Liu, and T. Jacks. 1997. p53-Independent apoptosis induced by paclitaxel through an indirect mechanism. Proc. Natl. Acad. Sci. USA 94:9679–9683. |
| 25. | Slowik, M. R., W. Min, T. Ardito, A. Karsan, M. Kashgarian, and J. S. Pober. 1997. Evidence that tumor necrosis factor triggers apoptosis in human endothelial cells by interleukin-1-converting enzyme-like protease-dependent and -independent pathways. Lab. Invest. 77:257–267. |
| 26. | Polunovsky, V. A., C. H. Wendt, D. H. Ingbar, M. S. Peterson, and P. B. Bitterman. 1994. Induction of endothelial cell apoptosis by TNF alpha: modulation by inhibitors of protein synthesis. Exp. Cell Res. 214:584–594. |
| 27. | Madge, L. A., and J. S. Pober. 2000. A phosphatidylinositol 3-kinase/Akt pathway, activated by tumor necrosis factor or interleukin-1, inhibits apoptosis but does not activate NFkappaB in human endothelial cells. J. Biol. Chem. 275:15458–15465. |
| 28. | Bannerman, D. D., M. Sathyamoorthy, and S. E. Goldblum. 1998. Bacterial lipopolysaccharide disrupts endothelial monolayer integrity and survival signaling events through caspase cleavage of adherens junction proteins. J. Biol. Chem. 273:35371–35380. |
| 29. | Goedert, M., M. Hasegawa, R. Jakes, S. Lawler, A. Cuenda, and P. Cohen. 1997. Phosphorylation of microtubule-associated protein tau by stress- activated protein kinases. FEBS Lett. 409:57–62. |
| 30. | Parker, C. G., J. Hunt, K. Diener, M. McGinley, B. Soriano, G. A. Keesler, J. Bray, Z. Yao, X. S. Wang, T. Kohno, and H. S. Lichenstein. 1998. Identification of stathmin as a novel substrate for p38 delta. Biochem. Biophys. Res. Commun. 249:791–796. |
| 31. | De Vos, K., F. Severin, F. Van Herreweghe, K. Vancompernolle, V. Goossens, A. Hyman, and J. Grooten. 2000. Tumor necrosis factor induces hyperphosphorylation of kinesin light chain and inhibits kinesin-mediated transport of mitochondria. J. Cell Biol. 149:1207–1214. |
| 32. | Ferrero, E., M. R. Zocchi, E. Magni, M. C. Panzeri, F. Curnis, C. Rugarli, M. E. Ferrero, and A. Corti. 2001. Roles of tumor necrosis factor p55 and p75 receptors in TNF-alpha-induced vascular permeability. Am. J. Physiol. Cell Physiol. 281:C1173–C1179. |
| 33. | Kiemer, A. K., N. C. Weber, R. Furst, N. Bildner, S. Kulhanek-Heinze, and A. M. Vollmar. 2002. Inhibition of p38 MAPK activation via induction of MKP-1: atrial natriuretic peptide reduces TNF-alpha-induced actin polymerization and endothelial permeability. Circ. Res. 90:874–881. |
| 34. | Clauss, M., C. Sunderkotter, B. Sveinbjornsson, S. Hippenstiel, A. Willuweit, M. Marino, E. Haas, R. Seljelid, P. Scheurich, N. Suttorp, M. Grell, and W. Risau. 2001. A permissive role for tumor necrosis factor in vascular endothelial growth factor-induced vascular permeability. Blood 97:1321–1329. |
| 35. | Kevil, C. G., T. Oshima, and J. S. Alexander. 2001. The role of p38 MAP kinase in hydrogen peroxide mediated endothelial solute permeability. Endothelium 8:107–116. |
| 36. | Lavoie, J. N., H. Lambert, E. Hickey, L. A. Weber, and J. Landry. 1995. Modulation of cellular thermoresistance and actin filament stability accompanies phosphorylation-induced changes in the oligomeric structure of heat shock protein 27. Mol. Cell. Biol. 15:505–516. |
| 37. | Guay, J., H. Lambert, G. Gingras-Breton, J. N. Lavoie, J. Huot, and J. Landry. 1997. Regulation of actin filament dynamics by p38 MAP kinase-mediated phosphorylation of heat shock protein 27. J. Cell Sci. 110:357–368. |
| 38. | Huot, J., F. Houle, S. Rousseau, R. G. Deschesnes, G. M. Shah, and J. Landry. 1998. SAPK2/p38-dependent F-actin reorganization regulates early membrane blebbing during stress-induced apoptosis. J. Cell Biol. 143:1361–1373. |
| 39. | Rousseau, S., F. Houle, J. Landry, and J. Huot. 1997. p38 MAP kinase activation by vascular endothelial growth factor mediates actin reorganization and cell migration in human endothelial cells. Oncogene 15:2169–2177. |
