Microtubule disassembly increases endothelial cell barrier
dysfunction: role of MLC phosphorylation
Alexander D.
Verin,
Anna
Birukova,
Peiyi
Wang,
Feng
Liu,
Patrice
Becker,
Konstantin
Birukov, and
Joe G. N.
Garcia
Division of Pulmonary and Critical Care Medicine, Johns
Hopkins University School of Medicine, Baltimore, Maryland 21224
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ABSTRACT |
Endothelial cell (EC) barrier
regulation is critically dependent on cytoskeletal components
(microfilaments and microtubules). Because several edemagenic agents
induce actomyosin-driven EC contraction tightly linked to myosin light
chain (MLC) phosphorylation and microfilament reorganization, we
examined the role of microtubule components in bovine EC barrier
regulation. Nocodazole or vinblastine, inhibitors of microtubule
polymerization, significantly decreased transendothelial electrical
resistance in a dose-dependent manner, whereas pretreatment with the
microtubule stabilizer paclitaxel significantly attenuated this effect.
Decreases in transendothelial electrical resistance induced by
microtubule disruption correlated with increases in lung permeability
in isolated ferret lung preparations as well as with increases in EC
stress fiber content and MLC phosphorylation. The increases in MLC
phosphorylation were attributed to decreases in myosin-specific
phosphatase activity without significant increases in MLC kinase
activity and were attenuated by paclitaxel or by several strategies (C3
exotoxin, toxin B, Rho kinase inhibition) to inhibit Rho GTPase.
Together, these results suggest that microtubule disruption initiates
specific signaling pathways that cross talk with microfilament
networks, resulting in Rho-mediated EC contractility and barrier dysfunction.
transendothelial electrical resistance; nonmuscle contraction; actin rearrangement
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INTRODUCTION |
THE VASCULAR ENDOTHELIUM
ACTS as a selective barrier between the vascular space and
underlying tissues. Compromise of endothelial cell (EC) barrier
integrity leads to an increase in vascular permeability, a cardinal
feature of inflammation resulting in tissue edema, hypoxemia, and,
often, increased morbidity and mortality. Substantial work, including
work from our own laboratory (23), has verified that the EC barrier is regulated by contractile and tethering mechanisms, the effects of which are critically dependent on EC cytoskeletal components (microfilaments, intermediate filaments, and
microtubules). For example, our laboratory has previously shown
(21, 58) that specific edemagenic agents such as the serine protease thrombin induce EC barrier dysfunction, primarily via
actomyosin-driven contraction initiated by myosin light chain (MLC)
phosphorylation and tightly linked to microfilament reorganization. Thrombin increases Ca2+ and EC centripetal tension
(41, 46), and 20-kDa MLC
(MLC20) phosphorylation peaks at 2 min and returns
nearly to control levels by 60 min, indicating involvement of both MLC
kinase (MLCK) and MLC phosphatase (PPase) activity in the thrombin
response (21, 57). At least two responsible classes of
MLCKs are potentially able to phosphorylate MLC in vivo (4, 21,
29), including the newly cloned endothelial
Ca2+/calmodulin (CaM)-dependent MLCK (22) and
Rho kinase, the activity of which depends on activation of the small G
protein Rho.
It has become well established that Ras-related GTPases of the Rho
family organize the actin cytoskeleton and regulate focal adhesion formation. Rho GTPases are inactive in the GDP-bound form
and are activated by GDP/GTP exchange (17, 56). Rho can be
specifically inactivated by bacterial toxins by either ADP-ribosylation (Clostridium botulinum C3 exotoxin) or glucosylation
(Clostridium difficile, toxin B), which presumably block the
interaction of Rho with downstream targets like Rho kinase (1, 2,
50). There is a significant controversy, however, as to the
effect of Rho on EC contractile and barrier properties. For example, C3
exotoxin has been suggested to both attenuate EC thrombin-mediated contraction via reduction of the thrombin-induced decrease in myosin-specific PPase activity (20) and to disrupt EC
barrier properties, with minimal effect on EC contractile properties
(61). Our laboratory (26) has recently shown
that C3 exotoxin completely abolished thrombin- and diperoxovanadate
(DPV)-induced increases in MLC phosphorylation, suggesting the
involvement of Rho activation in the EC contraction produced by these
agonists. Downstream targets of the Rho/Rho kinase signaling cascade
include the regulatory subunit of myosin-specific PPase (M 130), the
phosphorylation of which by Rho kinase leads to dissociation from the
catalytic subunit, thereby decreasing myosin-specific PPase 1 activity
and subsequently enhancing the level of MLC phosphorylation (37, 39).
In contrast to the microfilamentous actin cytoskeleton, information
about the microtubule network and its linkage to the contractile processes is limited. Disassembly of microtubules caused rapid and
substantial strengthening of nonmuscle contractility and led to rapid
assembly of microfilament bundles and focal adhesions (7, 15,
28), an effect abolished by microtubule stabilization (15). Isometric contraction in fibroblasts induced by
microtubule disassembly correlated well with increases in MLC
phosphorylation (40); however, the molecular basis for
this finding has not been pursued. Furthermore, the role of the
microtubule network in EC shape changes, tension development, and
barrier properties is unknown. In this study, we examined the
biochemical mechanisms by which microtubule disassembly increases
endothelial contraction and barrier dysfunction and elucidated the
involvement of increased MLC phosphorylation in this process. These
studies confirm that MLC phosphorylation is a common mechanism by which
microtubule-disrupting agents and receptor-mediated agonists induce
nonmuscle contraction (40) and suggest an important
physiological role of the microtubule network in EC barrier regulation.
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METHODS |
Reagents.
EC cultures were maintained in medium 199 (GIBCO BRL, Chagrin Falls,
OH) supplemented with 20% (vol/vol) colostrum-free bovine serum
(Irvine Scientific, Santa Ana, CA), 15 µg/ml of EC growth supplement
(Collaborative Research, Bedford, MA), 1% antibiotic-antimycotic (10,000 U/ml of penicillin, 10 µg/ml of streptomycin, and 25 µg/ml of amphotericin B; K. C. Biologicals, Lenexa, KA), and 0.1 mM nonessential amino acids (GIBCO BRL). Unless specified, reagents were
obtained from Sigma (St. Louis, MO). PBS, Hanks' balanced salt solution
without phenol red, and LipofectAMINE were purchased from GIBCO BRL
(Grand Island, NY). DPV was kindly provided by Dr. V. Natarajan (Johns
Hopkins University, Baltimore, MD). C3 exotoxin, vinblastine, and ML-7
were purchased from Calbiochem (La Jolla, CA). Toxin B was purchased
from List Laboratories (Campbell, CA). MLC antibody was produced in
rabbit against baculovirus-expressed and purified smooth muscle MLC by
Biodesign International (Kennebunk, ME). Texas Red-phalloidin was
purchased from Molecular Probes (Eugene, OR). Anti-MLCK D119 antibody
was kindly provided by Dr. P. Gallagher (Indiana University,
Indianapolis, IN).
Bovine pulmonary artery EC culture.
ECs were obtained frozen at passage 16 from the American
Type Culture Collection (CCL 209; Manassas, VA) and were used between passages 19 and 24 as our laboratory has
previously described in detail (21, 59).
Endothelial monolayer resistance determinations.
The electrical resistance of EC monolayers was measured with the
electrical cell impedance sensor technique as previously described
(24, 55). In this system (Applied Biophysics, Troy, NY),
the cells are cultured on a small gold electrode (10
4
cm2) in DMEM (GIBCO BRL) supplemented with 20% (vol/vol)
colostrum-free bovine serum, antibiotics, and growth factors as
described in Reagents. The cells act 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 was applied to the ECs through a 1-M
resistor,
creating an approximate constant current source (1 µA). The lock-in
amplifier attached to the electrodes detected changes in both magnitude
and phase of the voltage appearing across the ECs and was controlled by
an IBM-compatible personal computer that was used both to run the
experiments and process the data. Electrical resistance increased
immediately after the cells attached to and covered the electrodes, and
the resistance achieved a steady state when the ECs became confluent.
Thus experiments were conducted after the electrical resistance
achieved a steady state. Resistance data were normalized to the initial
voltage and plotted as a normalized resistance.
Isolated perfused ferret lung preparation.
Commercially available ferrets were anesthetized with pentobarbital
sodium (50 mg/kg ip), ventilated (20 breaths/min, tidal volume 12 ml/kg) via tracheostomy with warmed, humidified gas containing 40%
O2, and then rapidly exsanguinated as previously described
(6). After exsanguination, the ventilatory gas mixture was
switched to 16% O2-5% CO2, the ventilatory
rate was adjusted to 10 breaths/min, and an end-expiratory pressure of
3 mmHg was added. Lungs were isolated by insertion of cannulas into the
left atrium via the left ventricle and the pulmonary artery via the right ventricle, and then residual blood was flushed from the lungs
with physiological salt solution (PSS) containing 5 mM dextrose, 3 g/dl
of porcine albumin, and 2 g/dl of Ficoll (6). Isolated lungs were perfused at constant flow (50 ml · kg
1 · min
1) for 60 min
with PSS-0.1% ethanol (control) or PSS containing 1 µM vinblastine.
Pulmonary arterial, left atrial, and airway pressures were continuously
monitored (Grass model 7) with Statham P50 transducers referenced to
the left atrium. Glucose concentration and pH were monitored throughout
the perfusion period and did not differ among preparations.
After 60 min of extracorporeal perfusion, the pulmonary vasculature was
filled with PSS containing washed ferret red blood cells (hematocrit
20%), and the pulmonary arterial and left atrial cannulas were
connected to a common reservoir containing the same solution. The
reservoir was pressurized to 30 mmHg for 20 min, and samples were
rapidly withdrawn from the left atrial cannula for measurement of red
blood cell and albumin concentrations as previously described
(6). The osmotic reflection coefficient for albumin
(
alb) was calculated iteratively from the rate of change
of albumin concentration relative to red blood cell concentration (6).
MLC phosphorylation.
This assay was performed as our laboratory has previously described in
detail (21, 55).
Cytotoxicity assay.
A cytotoxicity assay was performed using a LIVE/DEAD
viability/cytotoxicity kit (Molecular Probes) according to the
manufacturer's protocol. This kit provides a two-color fluorescence
cell viability assay that is based on the simultaneous dual
determination of two recognized parameters of cell viability:
intracellular esterase activity (calcein) and plasma membrane integrity
(ethidium homodimer-1). Calcein produces an intense green fluorescence
in viable cells, whereas ethidium homodimer-1 enters cells with damaged
membranes and binds to nucleic acids, thereby producing a bright red
fluorescence in nonviable cells.
Permeabilization of ECs.
It is well known that C. botulinum C3 exoenzyme does not
easily pass through the cell membrane under native conditions. To penetrate the cell membrane, EC monolayers (80-100% confluence) grown on 60-mm culture dishes were rinsed with OptiMEM-I medium and
LipofectAMINE reagent (GIBCO BRL) added at a final concentration of 20 µg/ml for 1 h followed by the addition of C3 exoenzyme (2.5 µg/ml) for an additional 11 h as we have previously described in
detail (9).
Preparation of silicone rubber substrates.
To visualize cell contractility, flexible rubber substrates were
generated as previously described (8, 31). Approximately 15 µl of silicone monomer (dimethyldiphenylpolysiloxane; Sigma) were
applied to 18-mm glass coverslips and allowed to spread for 30 min.
Cells were plated on the top of thin polymerized flexible rubber
silicone films, and contractility was assessed by formation of wrinkles
on the silicone substrate as observed with time-lapse videomicroscopy.
Western immunoblotting.
Protein extracts were separated by SDS-PAGE, transferred to
nitrocellulose or polyvinylidene difluoride membranes (30 V for 18 h or 90 V for 2 h), and reacted with specific antibodies of interest. Immunoreactive proteins were detected with the enhanced chemiluminescent detection system (ECL) according to the
manufacturer's directions (Amersham, Little Chalfont, UK). The
relative intensities of the protein in the bands were quantified by
scanning densitometry.
Determination of myosin-specific and total serine/threonine PPase
activities in endothelium.
ECs from D100-mm dishes were treated with vehicle (0.1%
methanol or DMSO), 0.2 µM vinblastine, or 10 nM calyculin for 10, 30, or 60 min. To prepare total cell lysates, the cells were rinsed twice with ice-cold Tris-buffered saline (20 mM Tris · HCl, pH 7.6, and 137 mM NaCl) and buffer A (50 mM
Tris · HCl, pH 7.0, 0.1 mM EDTA, and 28 mM mercaptoethanol);
then 400 µl of buffer A were added to the dishes, and the
cells were quickly frozen at
70°C, scraped, and homogenized by
passing the cell suspension several times through a 1-ml tuberculin
syringe. Myosin-enriched fractions were prepared with 500 µl of total
cell lysate as our laboratory has previously described
(59). PPase activity was determined in a final volume of
20 µl with a Malachite Green (Upstate Biotechnology) 96-well plate
assay as previously described (18, 30). Total cell lysate
or myosin-enriched fraction (1 µl) was incubated with 0.25 mM
phosphopeptide KRpTIRR (specific serine/threonine PPase substrate;
Upstate Biotechnology, Lake Placid, NY) in PPase assay buffer (50 mM
Tris · HCl, pH 7.0, 0.1 mM EDTA, 28 mM mercaptoethanol, and 0.1 mg/ml of BSA) at 30°C for 20 min. As a control, the cell suspension
and phosphopeptide were incubated in a PPase assay buffer containing
serine/threonine PPase inhibitors (1 mM EGTA and 5 µM okadaic acid).
Reactions were terminated by the addition of 100 µl of Malachite
Green solution, and the plates were incubated at room temperature for
15 min before measurement of absorbance at 620 nm. The phosphate
released in the enzyme reaction was determined by comparing the
absorbance over control value with the phosphate standard curve.
Reactions were carried out in triplicate. To ensure linear rates of
dephosphorylation, the extent of dephosphorylation was restricted
to <25%. The number of proteins contained in cell lysate was assessed
by TCA precipitation and then with a BCA protein assay kit (Pierce,
Rockford, IL) used according to the manufacturer's protocol. Basal
PPase activity in cell lysates was ~1,500 pmol phosphate · µg total
protein
1 · min
1.
Immunofluorescent microscopy.
ECs were grown to subconfluence on glass coverslips in DMEM. After
treatment, cells were fixed in 4% paraformaldehyde for 10 min at room
temperature. The cells were then washed three times with PBS,
permeabilized with 0.25% Triton X-100 in PBS for 5 min, and blocked
with 2% BSA in PBS for 30 min, and then actin was stained with Texas
Red-phalloidin (Molecular Probes) for 1 h at room temperature.
After three washes with PBS, the coverslips were mounted with a
SlowFade antifade kit (Molecular Probes). Analysis of the
stained cells was performed with a Nikon Eclipse TE 300 microscope
equipped with ×20 to ×100 objective lenses.
Determination of MLCK activity.
MLCK activity was determined in nondenaturing MLCK immunoprecipitates
with exogenous MLC as a substrate, as our laboratory previously
described in detail (58).
Statistics.
ANOVAs with a Student-Newman-Keuls test were used to compare the means
of kinase and PPase activities, the
alb, and the ratios of un-, mono-, and diphosphorylated MLCs of two or more different treatment groups. Results are expressed as means ± SE.
Differences between two groups were considered statistically
significant when P < 0.05.
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RESULTS |
Effect of microtubule alteration on EC barrier function.
To examine the effect of microtubule disruption on EC barrier function,
we monitored electrical resistance across EC monolayers [(transendothelial electrical resistance (TER)] after treatment with
the microtubule-disrupting agents nocodazole and vinblastine. Both
nocodazole (Fig. 1A) and
vinblastine (Fig. 1B) decreased TER in a dose-dependent
manner, with half-maximal declines in TER after nocodazole
(0.3-5.0 µM) occurring in ~15-30 min. Maximal vinblastine
effect was observed at 0.5-2.5 µM; however, incubation of cells
with even 200 nM of vinblastine significantly decreased electrical
resistance (Fig. 1B). Vinblastine-induced EC permeability was also confirmed in vivo with the use of isolated perfused ferret lungs (Table 1) in which the
alb estimated in the control lungs was 0.7, a mean value
similar to that previously obtained in uninjured isolated ferret lungs
(6). In contrast, after 60 min of perfusion with
vinblastine, the
alb decreased to 0.35 (Table 1) in
association with significant lung weight gain and consistent with
vinblastine-induced increased vascular permeability and EC barrier
dysfunction. Pulmonary arterial and pulmonary airway pressures remained
almost constant (7.0-8.5 and 5.5-6.0 mmHg, respectively)
through 60 min of perfusion in the vinblastine preparation and were
statistically indistinguishable from control values. Both nocodazole
(Fig. 2) and vinblastine (data not shown)
perturbed EC barrier properties without cytotoxicity. We next examined
the effect of microtubule stabilization on barrier dysfunction induced
by microtubule-disrupting agents. Figure
3A demonstrates that the
microtubule-stabilizing agent paclitaxel did not affect basal TER but
significantly attenuated the decrease in TER induced by nocodazole.
Furthermore, paclitaxel reversed the effect of the
microtubule-disrupting agents on TER, indicating that the effect of
microtubule disruption on EC barrier dysfunction is reversible (Fig.
3B).

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Fig. 1.
Effect of microtubule disassembly on transendothelial
electrical resistance (TER). Disruption of microtubule structure by
microtubule inhibitors led to endothelial cell (EC) barrier compromise.
A: TER was monitored for 2 h. Arrow, time at which
cells were treated with either vehicle (0.1% DMSO) or nocodazole in
complete medium. B: TER was monitored for 6 h. Cells
were treated with either vehicle (0.1% methanol) or vinblastine in
complete medium. Results are from representative experiments
(n = 3 each).
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Fig. 2.
Effect of nocodazole on cell viability. Shown are
immunofluorescent images (original magnification, ×40) of EC
monolayers (80% confluence) stained with either calcein AM (1 µM for
45 min; A-C) for intracellular esterase activity or
ethidium homodimer-1 (4 µM for 45 min; D-F)
for plasma membrane integrity as described in MATERIALS AND
METHODS. A and D: vehicle (0.1%
DMSO)-treated ECs; B and E: ECs treated with 2 µM nocodazole for 45 min; C and F: ECs treated
with 70% methanol for 30 min. Control and nocodazole-treated ECs show
intense calcein staining, indicating the presence of ubiquitous
intracellular esterase activity ("live" cells), whereas
methanol-treated cells show intense ethidium staining, a characteristic
of cells with damaged membranes ("dead" cells). Results are from a
representative experiment (n = 3).
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Fig. 3.
Effect of microtubule stabilization on decrease in TER
induced by microtubule inhibitors. EC monolayer resistance was
monitored for 4 h. A: ECs were pretreated with either
vehicle (0.1% DMSO) or paclitaxel (Pacli; 2.5 µM) for 75 min
followed by challenge with either vehicle or nocodazole (Noco; 2 µM).
B: ECs were first treated by either vehicle, vinblastine
(Vin; 0.3 µM), or nocodazole (2 µM) for 30 min and then
challenged with either vehicle or paclitaxel (10 µM). Results are
from representative experiments (n = 3). Microtubule
stabilization significantly attenuated and reversed the effect of
microtubule inhibitors on TER.
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Effect of microtubule disassembly on EC microfilament
reorganization.
Because our laboratory (23, 25) and others,
(47) have demonstrated that agonist-induced EC barrier
dysfunction is critically dependent on cytoskeletal changes, we next
examined the effect of filamentous actin (F-actin) stabilization by
phalloidin on vinblastine-induced EC barrier dysfunction (Fig.
4). Phalloidin significantly
attenuated the decrease in electrical resistance induced by
vinblastine, indicating an important role for the actin cytoskeleton in
EC activation stimulated by microtubule disruption. To extend these
findings, we next studied the direct effect of nocodazole on the EC
actin cytoskeleton. Figure 5 shows
immunofluorescent images of EC confluent monolayers stained for F-actin
with Texas Red-phalloidin. As seen in Fig. 5A, control cells
have F-actin organized primarily in a cortical ring, with few
actin-associated stress fibers spanning the cell. Nocodazole produces
dramatic cortical actin dissolution and a significant increase in
stress fibers (Fig. 5B), reflecting contraction. Stress
fiber formation induced by nocodazole leads to gap formation,
reflecting barrier compromise (Fig. 5B).

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Fig. 4.
Effect of microfilament stabilization on
vinblastine-induced EC barrier dysfunction. ECs grown on gold
microelectrodes were pretreated with the F-actin stabilizer phalloidin
(Pha; 1 µM) for 60 min followed by vinblastine (0.3 µM). Arrows,
time of addition. TER was assessed as described in MATERIALS AND
METHODS. Results are from representative experiments
(n = 3). Stabilization of actin filaments significantly
attenuated the decrease in TER induced by microtubule inhibitors.
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Fig. 5.
Effect of microtubule alteration on F-actin rearrangement.
Immunofluorescent images (original magnification, ×60) show confluent
ECs stained for F-actin with Texas Red-phalloidin. ECs were treated
with either vehicle (0.1% DMSO; A) or nocodazole
(B) for 30 min. Disruption of microtubules led to
significant increases in stress fiber formation, indicating EC
contraction with development of intercellular gaps (arrows) indicating
barrier compromise.
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Effect of microtubule disruption on MLC phosphorylation.
Our laboratory (21) and others (66) have
previously shown the critical involvement of MLC phosphorylation in
agonist-induced barrier dysfunction in macro- and microvascular
endothelium. Similar to DPV and thrombin treatments, both of which
increase MLCK activity and MLC phosphorylation in ECs (22, 26,
57, 58), pretreatment of ECs with microtubule inhibitors
produced significant enhancement of MLC phosphorylation in a
time-dependent manner (Fig.
6A). The increases in MLC
phosphorylation produced by microtubule disruption (up to 1 mol
phosphate/mol MLC, peak at 30-60 min) were less than the levels of
DPV-induced MLC phosphorylation (up to 1.8 mol/mol MLC)
(26) but similar to the level of thrombin-induced
MLC phosphorylation (up to 1.2 mol/mol MLC) (21). Neither
vinblastine nor nocodazole at lower (up to 100 nM) concentrations had
significant effects on the level of MLC phosphorylation (data not
shown). The increased MLC phosphorylation produced by microtubule
inhibitors was completely abolished by microtubule stabilization with
paclitaxel (Fig. 6B). Increases in MLC phosphorylation
correlated well with the time frame of decreased TER produced by
microtubule assembly inhibition (shown in Fig. 1), suggesting the
involvement of contractile mechanisms in EC barrier dysfunction induced
by microtubule disruption. Cell contraction induced by microtubule
disruption was confirmed by an increase in the number of wrinkles
produced by EC monolayers grown on a thin silicone film (Fig.
7).

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Fig. 6.
Effect of microtubule disruption on EC myosin light chain
(MLC) phosphorylation. A: EC monolayers were pretreated with
either 0.3 µM vinblastine (top), 2 µM nocodazole
(bottom), or 5 µM diperoxovanadate (DPV; positive
control; top and bottom) for indicated times. MLC
phosphorylation was monitored by urea gel electrophoresis followed by
Western immunoblotting with anti-MLC antibody as previously described
by our laboratory (21, 55). un-P, unphosphorylated;
mono-P, monophosphorylated; di-P, diphosphorylated. B: EC
monolayers were pretreated by either vehicle (0.1% DMSO) or 10 µM
paclitaxel for 60 min and then treated with vehicle or 2 µM
nocodazole for 30 min, and the level of MLC phosphorylation was
quantitated by densitometry of Western blots probed with anti-MLC
antibody. Both microtubule inhibitors significantly increased MLC
phosphorylation in a time-dependent manner, with maximal effect at
30-60 min. The increased MLC phosphorylation induced by
microtubule disruption was completely abolished by microtubule
stabilization.
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Fig. 7.
Effect of nocodazole on EC contraction. Shown are phase-
contrast images of living cells (original magnification, ×20) grown on
silicone membranes as described in MATERIALS AND METHODS.
A: ECs before stimulation. B: same field after
treatment with nocodazole (2 µM) for 30 min. Note numerous
wrinkles formed by focally contracting ECs on the silicone substrate
(arrows).
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Effect of microtubule disruption on MLCK and myosin PPase
activities.
Because MLC phosphorylation status is determined by the balance between
MLCK and myosin PPase activities, we next determined the effect of
microtubule disruption on EC MLCK activity measured in the MLCK
immunoprecipitates in the presence of optimal Ca2+/CaM
availability (maximal activity) in either the presence or absence of
the specific MLCK inhibitor ML-7 or the Ca2+ chelator EGTA.
The data in Fig. 8A
demonstrate that inhibition of MLCK with ML-7 and EGTA completely
abolished basal MLCK activity, confirming that MLCK activity assay is
valid; however, neither form of microtubule inhibition altered MLCK
activity above basal levels in the presence of Ca2+/CaM.
Consistent with these results, ML-7 also failed to alter the MLC
phosphorylation induced by nocodazole, whereas thrombin-induced MLC
phosphorylation was abolished (Fig. 8B).

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Fig. 8.
Effect of microtubule disruption on MLC kinase (MLCK)
activity. A: confluent EC monolayers were pretreated with
either vehicle (0.1% DMSO), 2 µM nocodazole, or 0.2 µM vinblastine
for 30 min, and MLCK activity was assessed in nondenaturing MLCK
immunoprecipitates. Purified MLC was used as the preferred substrate in
the presence of either 0.3 mM CaCl2/1 µM calmodulin
(Ca2+/CaM; maximal activity), the specific MLCK inhibitor
ML-7 (10 µM), or the Ca2+ chelator EGTA (2 mM) as
described in MATERIALS AND METHODS. Data are means ± SE; n = 3 experiments. B: ECs were
pretreated with either vehicle (0.1% DMSO) or ML-7 (25 µM for 40 min) and then treated with thrombin (100 nM for 2 min) or nocodazole
(2 µM for 30 min), and MLC phosphorylation was assessed by urea
gel electrophoresis. Microtubule disruption had no effect on MLCK
activity in endothelium. +, Presence; , absence.
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To examine the involvement of protein PPases in the increase of MLC
phosphorylation induced by microtubule inhibitors, we next determined
the effect of vinblastine and calyculin (a potent serine/threonine
PPase inhibitor) (33, 59) on PPase activity in cell
homogenates (Fig. 9A) and the
myosin-specific PPase activity (59) present in the
myosin-enriched fraction (Fig. 9B). In contrast to
calyculin, vinblastine did not affect total PPase activity but
significantly decreased myosin-specific PPase activity, suggesting an
involvement of myosin PPase inhibition in the increased levels of MLC
phosphorylation induced by microtubule disruption.

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Fig. 9.
Effect of vinblastine and calyculin on serine/threonine
phosphatase (PPase) activity in endothelium. Confluent ECs from 100-mm
dishes were treated with either vehicle (0.1% methanol or 0.1% DMSO),
vinblastine (0.2 µM), or calyculin (10 nM) for indicated times. Cells
were then lysed, and the myosin-enriched fraction was prepared as
described in MATERIALS AND METHODS. Serine/threonine PPase
activity of both total cell lysates (A) and myosin-enriched
cell fractions (B) was measured with a serine/threonine
PPase assay kit. Microtubule disruption led to significant decreases in
myosin-specific PPase activity but not in total PPase activity in
endothelium. Data are means ± SE normalized as percent control
for each independent experiment; n = 4. *Significant
difference from control.
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Involvement of Rho in EC barrier dysfunction induced by microtubule
disruption.
In smooth muscle and nonmuscle cells including endothelium, myosin
PPase activity is mediated by activation of the small GTP-bound protein
Rho (20, 39). To examine the role of Rho in MLC
phosphorylation induced by microtubule disruption, we first used the
specific Rho inhibitor C3 exotoxin, which ADP-ribosylates and thereby
inactivates Rho (2, 3). Because C3 exotoxin is not cell
permeable, we initially permeabilized EC monolayers with LipofectAMINE
to enhance C3 exotoxin access and found that the toxin completely
abolished increases in MLC phosphorylation induced by either thrombin
or vinblastine (Fig. 10A),
indicating Rho involvement. Consistent with these data, the
cell-permeable toxin B, which glucosylates and inactivates small Rho
family G proteins (Rho, CDC 42, and Rac) (2, 13, 36), also
completely abolished nocodazole-induced MLC phosphorylation (Fig.
10B). In addition, inhibition of the Rho downstream target
Rho kinase by the specific cell-permeant inhibitor Y-27632
significantly attenuated nocodazole-induced decreases in TER (Fig.
11) and increases in stress fibers
(data not shown) and completely abolished nocodazole-induced
MLC phosphorylation (Fig. 11, inset), demonstrating the
direct involvement of Rho kinase in nocodazole-induced MLC
phosphorylation and permeability.

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Fig. 10.
Effect of Rho inhibition on increase in MLC
phosphorylation induced by microtubule disruption. Representative
Western blots (n = 3) of MLC separated by urea gel
electrophoresis are shown. A: ECs were first permeabilized
by LipofectAMINE treatment in the presence and absence of the specific
Rho inhibitor C3 exotoxin (2) and then treated with either
100 nM thrombin (Thr) for 2 min or 0.3 µM vinblastine for indicated
time periods. B: ECs were pretreated with either DMEM
(control) or the cell-permeable inhibitor of Rho family proteins toxin
B (TB; 1 µM) for 60 min and then treated with either vehicle (0.1%
DMSO) or nocodazole (2 µM) for 30 min. Rho inhibition completely
abolished the increase in MLC phosphorylation induced by microtubule
disruption.
|
|

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|
Fig. 11.
Effect of Rho kinase inhibition on nocodazole-induced decrease in
TER. TER was monitored for 5 h. At time indicated by
left arrowhead, ECs were pretreated with either vehicle
(0.05% DMSO) or the specific Rho kinase inhibitor Y-27632 (10 µM)
followed by challenge (right arrowhead) with either vehicle
(0.05% DMSO) or nocodazole (2 µM). Inhibition of Rho kinase by
Y-27632 significantly attenuated but did not completely abolish the
nocodazole-induced decrease in resistance of EC monolayers, consistent
with the primary role for activation of Rho pathway in
nocodazole-induced EC barrier dysfunction. Results from a
representative experiment (n = 3) are shown.
Inset: in the same experiment, Rho kinase inhibition
completely abolished nocodazole-induced MLC phosphorylation.
|
|
 |
DISCUSSION |
In this study, we have attempted to clarify the biochemical
pathway by which changes in EC permeability are linked to microtubule assembly and disassembly. We used primarily a pharmacological approach
to either disrupt or preserve microtubule organization (34) with agents such as nocodazole, a synthetic
antitubulin agent that reversibly blocks the self-assembly of tubulin
and depolymerizes preformed microtubules, and vinblastine, a vinca alkaloid that rapidly and reversibly binds to tubulin causing microtubule depolymerization and inhibition of microtubule assembly. Paclitaxel (taxol), a potent microtubule assembly-promoting and -stabilizing agent, prevented and reversed the effect of microtubule disruptors on tubulin assembly. We observed that both nocodazole and
vinblastine caused significant dose-dependent decreases in TER in
bovine pulmonary artery ECs, indicating EC barrier dysfunction. Stabilization of the microtubule network by paclitaxel significantly attenuated and reversed the decline in electrical resistance induced by
microtubule disruption. Consistent with our results, microtubule disassembly has been noted to decrease electrical resistance across thyroid epithelial cells (65), increase microvascular
permeability of the rat small intestine vasculature (51),
and increase the permeability and rate of monocyte transendothelial
migration across human macro- and microvascular ECs (38,
62). We further demonstrated that EC barrier dysfunction induced
by microtubule disruption is linked to decreases in cortical actin and
increases in stress fiber formation and endothelial contraction as
confirmed by deformation of a silicone substrate (i.e., wrinkling).
Microtubule-mediated contractility has recently been demonstrated in
smooth muscle (5, 43, 52, 54). In nonmuscle cells such as
fibroblasts, microtubule disruption appeared to activate actin
polymerization and induce stress fibers and focal adhesions (7,
15, 16, 19, 35, 45). Cytoskeletal changes were accompanied by
contraction as evidenced by either the measurement of isometric force
generated from cells cultured within a collagen lattice
(40) or deformation of a silicone substrate
(15, 16).
The biochemical events linking the cytoskeletal changes initiated by
microtubule disruption and nonmuscle contraction are uncertain but
likely involve increased MLC phosphorylation and activation of the
small GTP-binding protein Rho (40, 45). Our laboratory
(21, 58) and others (29, 41, 48, 63, 64, 66)
have previously shown a direct link between the level of MLC
phosphorylation, EC retraction, and agonist-induced increases in EC
permeability, indicating the importance of actomyosin-driven contraction in EC barrier dysfunction. Our present data demonstrate for
the first time a correlation between microtubule disruption, activation
of the contractile machinery (MLC phosphorylation, stress fiber
formation), and a decrease in TER (barrier dysfunction). An increase in
MLC phosphorylation after microtubule disruption in ECs (present
study), previously noted in fibroblasts (40), suggests
activation of biochemical cascades that lead to contraction and EC
barrier failure. Importantly, fibroblasts precontracted with serum and
having increased levels of MLC phosphorylation showed a decreased
contractile response to microtubule disruption (40). This observation contradicts the
"tensegrity" model, which suggests that microtubules function as
compressive elements that oppose cellular contraction (10, 11,
32), and supports the active involvement of MLC phosphorylation
and Rho activation in the initiation of contraction after
microtubule disruption (16, 40).
Our laboratory (21, 57, 58) has previously shown that
thrombin-mediated EC cytoskeletal rearrangement and barrier dysfunction are strongly correlated with increased Ca2+/CaM-dependent
MLCK activity, myosin PPase inhibition, and a net increase in MLC
phosphorylation. In contrast to thrombin, microtubule inhibitors did
not increase MLCK activity, and specific inhibition of MLCK (with ML-7)
did not affect nocodazole-induced MLC phosphorylation, indicating that
MLCK is not involved in the activation of the EC contractile machinery
that is induced by microtubule inhibition. Increases in MLC
phosphorylation and the initiation of contraction in smooth muscle and
nonmuscle cells also follow the activation of the small GTPase Rho
(14, 53) and its target Rho kinase, which directly
phosphorylates MLC in vitro and in vivo (4, 42, 44). In
addition, Rho kinase phosphorylates the regulatory subunit of
myosin-associated PPase, which leads to inhibition of myosin-associated
PPase activity and an increase in MLC phosphorylation (37, 39,
49). Our laboratory (26) and others (12, 20, 27) have recently shown the involvement of Rho in
agonist-induced EC barrier dysfunction, increases in MLC
phosphorylation, and myosin PPase inhibition. Our present data indicate
that specific Rho or Rho kinase inhibitors completely abolish increases
in MLC phosphorylation induced by microtubule inhibitors, strongly
implicating the involvement of the Rho pathway in the EC barrier
dysfunction induced by microtubule disruption. Consistent with these
data, Rho kinase inhibition significantly attenuated nocodazole-induced decreases in TER. Although microtubule depolymerization in fibroblasts also induces stress fiber formation via Rho activation
(45), the precise mechanisms of Rho activation induced by
microtubule disruption are not currently known. Our laboratory (Verin
and Garcia, unpublished observations) demonstrated that Rho
inhibition did not directly affect EC microtubule structure, indicating
that Rho activation is a downstream event of microtubule disruption. To
evaluate biochemical events leading to Rho-dependent increases in MLC
phosphorylation and EC cytoskeletal rearrangement, we measured PPase
activity in the myosin-enriched fraction, which includes myosin, actin,
and regulatory and catalytic subunits of endothelial myosin PPase
(57, 59, 60), and most likely represents myosin-specific PPase activity in endothelium. We found that the microtubule inhibitor vinblastine significantly decreased myosin-specific, but not total, PPase activity in bovine ECs, supporting the involvement of myosin PPase in the increased MLC phosphorylation, cytoskeletal rearrangement, and barrier dysfunction induced by microtubule disruption.
In summary, we have characterized the biochemical and physiological
pathways leading to EC barrier dysfunction induced by microtubule
disruption. Microtubule disassembly leads to increased MLC
phosphorylation, stress fiber formation, and EC contraction via Rho
GTPase-dependent, but not MLCK-dependent, mechanisms that are likely
mediated via myosin PPase inhibition. Although the precise mechanism
that links microtubules, Rho activation, and EC barrier dysfunction
remains to be determined, our data demonstrate a significant role of
microtubule dynamics in EC barrier regulation.
 |
ACKNOWLEDGEMENTS |
We gratefully acknowledge Lakshmi Natarajan and Mary Ann Booth for
superb technical assistance. Special appreciation is extended to Dr. V. Natarajan (Johns Hopkins University, Baltimore, MD) and Dr. P. Gallagher (Indiana University, Indianapolis, IN) for providing key
reagents for this work.
 |
FOOTNOTES |
This work was supported by National Heart Lung, and Blood Institute
Grants HL-44746, HL-50533, HL-58064, and HL-67307 and funds from the
American Heart Association.
Address for reprint requests and other correspondence: A. D. Verin, The Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Circle, Baltimore, MD 21224 (E-mail:
averin{at}welch.jhu.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 22 November 2000; accepted in final form 29 March 2001.
 |
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