Role of
Ca2+/calmodulin-dependent
phosphatase 2B in thrombin-induced endothelial cell contractile
responses
Alexander D.
Verin,
Clare
Cooke,
Maria
Herenyiova,
Carolyn E.
Patterson, and
Joe G. N.
Garcia
Departments of Medicine, Physiology, and Biophysics, Indiana
University School of Medicine, Richard L. Roudebush Veterans
Administration Medical Center, Indianapolis, Indiana 46202
 |
ABSTRACT |
Thrombin-induced
Ca2+ mobilization, activation of
Ca2+/calmodulin-dependent myosin
light chain (MLC) kinase (MLCK), and increased phosphorylation of MLCs
precede and are critical to endothelial cell (EC) barrier dysfunction.
Net MLC dephosphorylation after thrombin is nearly complete by 60 min
and involves type 1 phosphatase (PPase 1) activity. We now report that
thrombin does not alter total PPase 1 activity in EC homogenates but
rather decreases myosin-associated PPase 1 activity. The PPase 1 inhibitor calyculin fails to prevent thrombin-induced MLC
dephosphorylation. However, thrombin significantly increased the
activity of Ca2+-dependent PPase
2B in EC homogenates (~1.5- to 2-fold), with PPase 2B activation
correlating with phosphorylation of the PPase 2B catalytic subunit.
Western immunoblotting revealed PPase 2B to be present in cytoskeletal
EC fractions, with specific PPase 2B inhibitors such as cyclosporin
(200 nM) and deltamethrin (100 nM to 1 µM) attenuating
thrombin-induced cytoskeletal protein dephosphorylation, including EC
MLC dephosphorylation. These results suggest a model whereby
thrombin-inducible contraction is determined by the phosphorylation
status of EC MLC regulated by the balance between EC MLCK, PPase 1 (constitutive), and PPase 2B (inducible) activities.
bovine pulmonary artery endothelium; thrombin-stimulated
endothelial cell permeability; thrombin-stimulated myosin light chain
phosphorylation; selective phosphatase 2B inhibitors
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INTRODUCTION |
A MAJOR FUNCTION of the vascular endothelial cell (EC)
monolayer is to serve as a semiselective barrier to fluid and solute flux across the blood vessel wall. The integrity of the EC monolayer is
an essential requirement for maintenance of this cellular barrier. Increased endothelial permeability is a prominent feature of acute lung
injury and inflammatory lung syndromes and is the result of
intercellular gap formation evoked by bioactive agents such as the
coagulation protease thrombin. Garcia and Schaphorst (19) and others (34) have employed a working model whereby EC gap formation
and barrier function is under close regulation by competing contractile
and tethering forces. Isometric force development appears to be
controlled, at least in part, by actin-myosin interaction regulated by
the phosphorylation status of the 20-kDa myosin "regulatory" light chain (MLC20). Garcia et
al. (16) have shown that thrombin-induced EC gap formation and barrier
dysfunction require protein kinase (PK) C activation and are coupled to
myosin light chain (MLC) phosphorylation catalyzed by a
high-molecular-mass
Ca2+/calmodulin (CaM)-dependent
MLC kinase (MLCK) isoform in endothelium. Inhibition of MLCK or
activation of cAMP-dependent PKA attenuated MLC phosphorylation and
prevented thrombin-induced increases in permeability (16). Changes in
the phosphorylation state of MLC are determined by the balance between
MLCK and myosin-specific phosphatase (PPase) activities (48). Verin et
al. (51) have previously shown that calyculin, a potent type 1 and 2A
PPase inhibitor, increased MLC phosphorylation and permeability of EC monolayers and produced gaps in intact and microinjected ECs, suggesting a potential role for PPases in the regulation of EC contractility and barrier function. Although several PPases are capable
of dephosphorylating isolated MLC, only type 1 myosin-associated PPase
was able to effectively dephosphorylate native smooth and skeletal
muscle myosin (1, 7). Type 2A PPase was effective toward isolated MLC
but ineffective in dephosphorylation of native myosin (1, 7). Recently,
the implied involvement of a
Ca2+-regulated PPase (2B or
calcineurin) in the maintenance of catch tension via MLC
dephosphorylation in molluscan muscles (6, 26), as well as the tight
association of PPase 2B with the nonmuscle cytoskeleton (13, 38), has
raised the possibility that PPase 2B may also be involved in the
regulation of contractility in specific cell types. Although Verin et
al. (51) recently determined that a type 1 PPase is primarily
responsible for the physiological dephosphorylation of myosin in ECs,
myosin-specific PPase properties in the endothelium remain incompletely
understood. In the present study, we found that, similar to
constitutively active myosin-associated PPase 1, thrombin-inducible
Ca2+/CaM-dependent PPase 2B may be
also involved in agonist-mediated EC activation. Furthermore, in
contrast to smooth muscle contraction, thrombin-mediated EC
contraction-relaxation could require activation of a complex PPase
cascade consisting of PPases 1 and 2B.
 |
METHODS |
Materials. Rabbit
skeletal muscle glycogen phosphorylase and phosphorylase kinase were
purchased from GIBCO (Gaithersburg, MD). Histone type II-S from calf
thymus and cAMP-dependent PKA catalytic subunit were obtained from
Sigma (St. Louis, MO); CaM was purchased from Ocean Biologics (Edmonds,
WA). Purified PPases and polyclonal anti-PPase 1 and anti-PPase 2A
antibodies were purchased from Upstate Biotechnology (Lake Placid, NY).
Anti-PPase 2B (calcineurin) polyclonal antibody was purchased from
either Chemicon (Temecula, CA) or Transduction Laboratories (Lexington, KY). Anti-CS1
antibody was kindly provided by Dr. Anna De Paoli Roach (Indiana University, Indianapolis, IN) and MLC-specific antibody
by Drs. James Stull (University of Texas, Dallas, TX) and Susan Gunst
(Indiana University, Indianapolis, IN). Deltamethrin was
purchased from BIOMOL Research Laboratories (Plymouth Meeting, PA),
cyclosporin was obtained from Indiana University Pharmacy (Indianapolis, IN), and calyculin A was obtained from Sigma.
Nitrocellulose filters and kaleidoscope-prestained standards were
obtained from Bio-Rad Laboratories (Hercules, CA). Prestained
molecular-mass standards for SDS-PAGE were purchased from GIBCO (Grand
Island, NY). Enhanced chemiluminescence (ECL) developing kit was
purchased from Amersham (Little Chalfont, United Kingdom),
125I-labeled protein A was
purchased from ICN Radiochemicals (Irvine, CA), and
[
-32P]ATP was from
New England Nuclear (Wilmington, DE). Other reagents were reagent
grade and obtained from either Sigma or Bio-Rad.
Bovine pulmonary artery endothelial cell
culture. Bovine pulmonary artery endothelial cells
(BPAECs) were obtained frozen at 16 passages from American Type Culture
Collection (culture line CCL 209; Manassas, VA) and were utilized at
passages
19-24 as
previously described (16, 51). Cells were cultured in DMEM (GIBCO BRL)
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 solution (10,000 units/ml of penicillin, 10 µg/ml of streptomycin, and 25 µg/ml of amphotericin B; K. C. Biologicals, Lenexa, KS), and
0.1 mM nonessential amino acids (GIBCO) and maintained at 37°C in a
humidified atmosphere of 5%
CO2-95% air. The ECs grew to
contact-inhibited monolayers with the typical cobblestone morphology. Cells from each primary flask were detached with 0.05% trypsin-EDTA, resuspended in fresh culture medium, and passaged to 100-mm dishes for
phosphatase studies, to 60-mm dishes for
32P labeling, to polycarbonate
filters for permeability studies, or into
75-cm2 flasks for MLC
phosphorylation studies.
MLC phosphorylation in intact
endothelium. MLC20
phosphorylation profiles were analyzed by urea-PAGE as previously
described by Garcia et al. (16) and Verin et al. (51). Briefly,
confluent BPAECs in 75-cm2 tissue
flasks were incubated with or without phosphatase inhibitors and
thrombin. At specified times, the medium was removed, the cells were
harvested by scraping into 10% trichloroacetic acid (TCA)-10 mM
dithiothreitol (DTT). After centrifugation, the pellets were washed
three times with diethyl ether and suspended in 6.7 M urea sample
buffer, and ~75 µg protein/lane were run on a 10% polyacrylamide-40% glycerol gel to separate unphosphorylated MLC from
phosphorylated MLC, which migrates more rapidly. The proteins were then
transferred to nitrocellulose (25 V for 90 min), and both
phosphorylated and unphosphorylated
MLC20 were detected by immunostaining overnight with an
MLC20-specific antibody (1:2,000 dilution) followed by a 1-h treatment with peroxidase-conjugated secondary antibodies (1:3,000 dilution; Bio-Rad). The blot was scanned
on a Bio-Rad densitometer, and the percent maximal MLC phosphorylation
was expressed by dividing the sum of two times the diphosphorylated MLC
area plus the monophosphorylated MLC area by the total of
phosphorylated and unphosphorylated areas (i.e., maximum
phosphorylation would be 200%). This method is very reproducible
because it measures a ratio of peak areas and thus is independent of
sample loading.
Endothelial monolayer permeability
determination. Macromolecule permeability of EC
monolayers was performed as previously described by Patterson et al.
(40). Gelatinized polycarbonate micropore membranes (0.8-µm pore
size; Corning, Acton, MA) were mounted on the base of
plastic cylinders and sterilized with ultraviolet light for 24 h.
BPAECs (2 × 105 cells/well)
were then seeded on the membrane and grown to confluence. The system
consists of two compartments, the upper (luminal) and the lower
(abluminal), which are separated by the polycarbonate filter on which
the EC monolayer is grown. The lower compartment was stirred
continuously and kept at a constant temperature of 37°C by use of a
thermally regulated water bath. Medium 199 with 25 mM HEPES in lieu of
bicarbonate was used in both compartments. BSA (4% final
concentration) complexed to Evans blue dye (EB-BSA) was added to the
luminal compartment, and samples were taken from the abluminal
compartment every 10 min for 60 min to establish the basal albumin
clearance (baseline) and then for an additional 60- to 120-min period
after each specific intervention. Transendothelial cell albumin
transport was determined by measuring the absorbance of Evans blue dye
in abluminal chamber samples at 620 nm in a spectrophotometer (Vmax
Multiplate Reader, Molecular Devices, Menlo Park, CA). Albumin
clearance rates were calculated by linear regression analysis for
control and experimental groups.
PPase activities in EC homogenates.
PPase 1 and 2A activities against phosphorylase A were determined as
previously described in detail by Verin et al. (51). To measure PPase
activity against 32P-labeled
histone II-S (Sigma), EC monolayers were washed two times with
phosphate-buffered saline (PBS; 10 mM phosphate buffer, 2.7 mM KCl, and
137 mM NaCl, pH 7.4; Sigma) and two times with ice-cold homogenization
buffer (50 mM HEPES and 28 mM
-mercaptoethanol, pH 7.4) containing
proteinase inhibitors [0.5 mM phenylmethylsulfonyl fluoride, 0.1 mM
N
-p-tosyl-L-lysine
chloromethyl ketone (TLCK), 0.1 mM leupeptin, and 2 mM
benzamidine]. Homogenization buffer (200 µl) was added to the
EC monolayers, and the plates were scraped and homogenized by passing
the cell suspension several times through a 1-ml tuberculin syringe.
Aliquots of EC homogenates were diluted two times in assay buffer (50 mM HEPES, pH 7.4, containing 0.6 mM
CaCl2, 50 nM CaM, 28 mM
-mercaptoethanol, and 2 mg/ml of BSA). The diluted homogenates (50 µl) containing PPase activity were added to a reaction mixture
consisting of 25 µl of
32P-labeled histone [6
mg/ml; 6,000-9,000
counts · min
1
(cpm) · pmol
1]
in HEPES buffer, pH 7.4, containing 2.8 mM
-mercaptoethanol. For
measuring calyculin-insensitive PPase 2B activity, the assay buffer was
supplemented with 30 nM calyculin A. After 30 min at 30°C, the
reaction was terminated by the addition of 15 µl of 100%
(wt/vol) ice-cold TCA. After 15 min on ice, the suspension was
centrifuged for 5 min (IEC Centra-M centrifuge, International Equipment), and aliquots (45 µl for each sample) of supernatant were
loaded onto Whatman 3 MM paper, dried for 15 min, and counted in
Scintiverse scintillation solution (Fisher Scientific, Fair Lawn, NJ)
by beta scintigraphy (Beckman model LS 6000 IC). Reactions were carried
out in duplicate, and controls consisted of incubations in which the
PPase-containing cell preparation was replaced by assay buffer. To
ensure linear rates of dephosphorylation, the extent of
dephosphorylation was restricted to <25%. One unit of phosphatase
activity was defined as the amount of enzyme that released 1 nmol
32Pi/min.
Preparation of 32P-labeled substrates.
32P-labeled phosphorylase A
(1,000-3,000 cpm/pmol) was prepared by phosphorylation of
phosphorylase B with phosphorylase kinase as previously described in
detail by Verin et al. (51).
32P-labeled phosphohistone
(6,000-9,000 cpm/pmol) was prepared by phosphorylation of histone
type II-S (Sigma) with a catalytic subunit of porcine PKA (Sigma) as
previously described (11), with some modifications. Briefly, histone
(90 mg) was dissolved in 15 ml of 50 mM HEPES buffer, pH 7.4, including
10 mM magnesium acetate and 1 mM DTT. Next, PKA (30 µg) reconstituted
in 0.6 ml of water containing 40 mM DTT, 5 µM ATP, and 30 µCi of
[
-32P]ATP was added
to the histone solution, and the reaction mixture was incubated at
25°C for 1 h in a shaking water bath. The reaction was terminated
by adding 100% (wt/vol) TCA to a final concentration of 15%.
After centrifugation for 5 min at 10,000 g, the supernatant was discarded, and
the pellet was washed two times by resuspension and centrifugation in
15 ml of 15% ice-cold TCA. The pellet was dissolved in 7.5 ml of 0.1 M
NaOH and dialyzed first against 3 liters of PBS for 4 h and then
against the same volume of HEPES buffer, pH 7.4, containing 2.8 mM
-mercaptoethanol overnight. 32P-labeled substrates were stored
at 4°C.
EC detergent fractionation.
Myosin-enriched and myosin-depleted BPAEC fractions were prepared as
described previously in detail by Verin et al. (51). For preparation of
a cytoskeletal fraction enriched in actin, we used the procedure
described by Lee et al. (32), with some modifications. Briefly,
confluent BPAECs from 100-mm dishes were rinsed two times with 2 ml of
PBS (10 mM phosphate buffer, 2.7 mM KCl, and 137 mM NaCl, pH 7.4;
Sigma) at room temperature and incubated with 1.5 ml of extraction
buffer (1% Nonidet P-40, 150 mM NaCl, 50 mM NaF, and 28 mM
-mercaptoethanol in 50 mM Tris · HCl, pH 8.0)
containing proteinase inhibitors (0.5 mM phenylmethylsulfonyl fluoride,
0.1 mM TLCK, 0.1 mM leupeptin, and 2 mM benzamidine) for 30 min at
4°C. Extractable proteins were discarded, and the insoluble
proteins remaining on the dishes (i.e.,
detergent-insoluble cytoskeletal EC fractions) were rinsed
two times with ice-cold PBS, solubilized by scraping the dishes into 3 ml of Laemmli SDS sample buffer (31), and subjected to Western
immunoblotting analysis as described in Immunoblotting
analysis of BPAEC fractions. Protein
concentrations were determined by the Bradford (5) method with BSA as a
standard.
Phosphorylation of cytoskeletal
proteins. For the study of cytoskeletal protein
phosphorylation, the cells were initially loaded with
[32P]orthophosphate
(0.4 mCi/ml) in phosphate-free DMEM (Sigma) in the presence of 1%
serum for 4 h. Cytoskeletal fractions were prepared as described in
EC detergent
fractionation, subjected to SDS-PAGE (31),
and stained for proteins with Coomassie blue R-250. The stained gels
were dried and subjected to autoradiography with ECL Hyperfilm
(Amersham).
Immunoblotting analysis of BPAEC
fractions. Homogenates or BPAEC fractions were
subjected to SDS-PAGE (31) on 9% gels and either stained with
Coomassie blue R-250 or electrophoretically transferred to a
nitrocellulose membrane as previously described (50). After transfer
for 17-18 h at 30 V, the nitrocellulose membrane was blocked for 3 h in 5% nonfat dry milk in PBS, pH 7.4, containing 0.1% Tween 20 and
then incubated with anti-PPase 2A (2.5 µg/ml), anti-CS1 (1:1,000
dilution), or anti-PPase 2B (1: 1,000 dilution) antibodies for 1 h.
Immunoreactive proteins were detected by autoradiography after binding
of 125I-protein A to the primary
immunocomplex or using an ECL detection system according to the
manufacturer's directions (Amersham).
PPase 2B immunoprecipitation. For
immunoprecipitation under denaturing conditions, confluent EC
monolayers in 60-mm tissue culture dishes were labeled with
[32P]orthophosphate
(0.5 mCi/plate) for 2.5 h in phosphate-free DMEM (Sigma) without serum,
followed by stimulation with either vehicle alone or thrombin (100 nM;
various times). The stimuli were then removed, the monolayers were
rinsed two times with 2 ml of medium, further rinsed with 2 ml of PBS,
and scraped into 100 µl of SDS-denaturing stop solution (PBS, pH 7.4, 1 mM EDTA, 1 mM EGTA, 50 mM NaF, 10 mM sodium phosphate, 0.2 mM
orthovanadate, 1% SDS, and 14 mM
-mercaptoethanol). The homogenate
was prepared by passing the cell suspension several times through a
16-gauge needle. Homogenates were heat treated at 110° for 5 min,
diluted 1:10 with 900 µl of PBS, and incubated with 50 µl of 10%
Pansorbin suspension (Formalin-hardened and heat-killed Cowan 1 strain
Staphylococcus aureus cells;
Calbiochem, La Jolla, CA) for 30 min at room temperature. Samples were
clarified by microcentrifugation for 5 min, and the supernatants were
incubated with 5 µl of anti-PPase 2B antibodies (60 min at room
temperature or overnight at 4°C; Chemicon, Temecula, CA), then with
50 µl of 10% Pansorbin suspension for 60 min at room temperature.
Immunocomplexes were pelleted by microcentrifugation for 5 min, washed
3 × 1 ml with PBS, solubilized in 100 µl of boiled 2×
SDS-Laemmli sample buffer (31), and then separated from Pansorbin beads
by microcentrifugation and subjected to SDS electrophoresis (31). After
electrophoresis, the proteins were transferred to nitrocellulose
membranes, and 32P signals were
detected by autoradiography at
70°C. To identify the PPase
2B position, the membranes were subsequently stained with specific
PPase 2B antibodies. The relative intensities of the
32P-labeled PPase 2B were
quantified by scanning densitometry.
Immunofluorescence. The fluorescent
imaging of PPase 2B and MLC cell localization was performed on BPAEC
monolayers grown to confluence on glass coverslips. After treatment,
the cells were fixed by exchanging medium with 5% paraformaldehyde, 50 mM phosphate, 75 mM NaCl, and 25 mM Tris, pH 7, on ice for 10 min. The
cells were thoroughly rinsed with buffer containing 150 mM NaCl and 50 mM Tris, pH 7.6, and then permeabilized by a 3.5-min treatment with
0.2% Triton in rinse buffer. The cells were again rinsed three times
and incubated at room temperature for 1 h with 1% BSA in rinse buffer.
The fixed, permeabilized cells were incubated with both rabbit
anti-smooth muscle MLC antibody (1:50 in Tris-NaCl-1% albumin buffer;
kindly provided by Dr. Susan Gunst) and mouse anti-calcineurin antibody
(1:50; Transduction Laboratories, Lexington, KY) overnight at 4°C.
After being rinsed to remove unbound primary antibody, the cells were
incubated for 1 h at room temperature with 30 µg/ml of labeled
secondary antibodies (FITC-conjugated donkey anti-rabbit IgG, 1:50, and
lissamine rhodamine-conjugated donkey anti-mouse IgG antibody, 1:50;
Jackson, West Grove, PA). The cells were examined with a ×60 oil
objective with the Bio-Rad MRC 1024 confocal microscope and excitation
with Ar-Kr laser at 568-nm excitation/598-nm emission for rhodamine and
488-nm excitation/522-nm emission for FITC at a 3-µm aperture. Data
were collected for 7-17 planar sections at 0.5-µm intervals by
Bio-Rad LaserSharp acquisition software, processed by MetaMorph Imaging
software (Universal, West Chester, PA), and printed on a thermal dye
diffusion printer (Kodak, Rochester, NY).
 |
RESULTS |
Effect of thrombin on MLC-specific PPase activity in
intact endothelium. The dynamics of thrombin-induced
MLC phosphorylation-dephosphorylation in BPAECs were monitored by
urea-glycerol gel electrophoresis followed by Western blotting with an
MLC-specific antibody as previously described by Garcia et al. (16) and
Verin et al. (51). Figure 1 shows that MLCs
are constitutively phosphorylated, with a stoichiometry of ~0.4 mol
phosphate/mol MLC. Thrombin treatment (100 nM) causes a rapid increase
in MLC phosphorylation, with maximal phosphorylation at 2 min (~0.9
mol phosphate/mol MLC), which gradually decreased in a time-dependent
manner, with MLC dephosphorylation nearly returning to baseline by 60 min, with a stoichiometry of ~0.5 mol phosphate/mol MLC (Fig. 1).
Because these results implicated the involvement of PPases in the
regulation of thrombin-induced EC MLC phosphorylation, we studied the
effect of the PPase inhibitors calyculin A (Fig.
2A) and
cyclosporin (Fig. 2B) on the
phosphorylation status of MLC in thrombin-stimulated and nonstimulated
ECs. Intact EC monolayers were pretreated with vehicle or specific
PPase inhibitors for 1 h at 37°C and then stimulated with 100 nM
thrombin for 2 and 60 min. The levels of unphosphorylated and mono- and
diphosphorylated MLC species were compared both with the basal MLC
phosphorylation profile present in ECs and with the level of maximal
thrombin-activated MLC phosphorylation (Fig. 2). EC pretreatment with
10 nM calyculin A [equally selective for PPase 1 and PPase 2A;
IC50 = 0.3-0.4 nM (24)]
significantly increased the level of basal and maximal thrombin-induced
MLC phosphorylation but did not completely prevent thrombin-induced MLC
dephosphorylation (Fig. 2A). In
contrast, 200 nM cyclosporin [selective inhibitor of PPase 2B;
IC50 = 7 nM (14)] had no
effect on either the basal level or the maximal thrombin-induced MLC phosphorylation but significantly attenuated the thrombin-induced MLC
dephosphorylation at 60 min (Fig.
2B). Another specific PPase 2B
inhibitor, deltamethrin [0.1-1 µM;
IC50 = 0.03 nM (11)]
exhibited similar effects in stimulated and control cells (Fig.
3). These data indicate that the activation
of both PPase 1 and the
Ca2+/CaM-dependent PPase 2B may be
involved in thrombin-mediated MLC dephosphorylation.

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Fig. 1.
Time course of endothelial cell (EC) myosin light chain (MLC)
phosphorylation after thrombin activation. Confluent EC monolayers were
rinsed 2 times with medium 199 (M199) to remove serum and incubated in
M199 with 25 mM HEPES (M199H) in lieu of bicarbonate. Cells were
treated with vehicle (Control; M199H) or 100 nM thrombin for indicated
time periods, and MLC phosphorylation was monitored by urea gel
electrophoresis followed by immunoblotting with anti-MLC antibodies and
quantitated by scanning densitometry as described in
METHODS. Data are means ± SE from
14 independent experiments. * Significant difference compared
with maximal level of thrombin-induced MLC phosphorylation,
P < 0.05.
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Fig. 2.
Effect of phosphatase (PPase) inhibitors on MLC phosphorylation in EC
monolayers. Confluent bovine pulmonary artery endothelial cells
(BPAECs) were rinsed as described in Fig. 1, pretreated with either
vehicle [0.1% DMSO (A) or
M199H (B)] or specific PPase
inhibitors [10 nM calyculin A
(A) or 200 nM cyclosporin
(B)] for 1 h, and then
stimulated with 100 nM thrombin for 2 and 60 min. MLC phosphorylation
was quantitated as described in
METHODS. Data are means ± SE from
at least 6 independent experiments. Calyculin significantly increased
levels of basal and maximal thrombin-induced MLC phosphorylation but
did not completely prevent thrombin-induced MLC dephosphorylation,
whereas cyclosporin had no effect on levels of basal and maximal MLC
phosphorylation but partially blocked thrombin-induced MLC
dephosphorylation. Significant difference
(P < 0.05): * compared with
basal level of MLC phosphorylation; ** between thrombin-alone and
inhibitor-thrombin groups at 2 min; *** between thrombin groups
at 2 and 60 min (with and without inhibitor separately);
# between maximal level of
inhibitor-induced and inhibitor-thrombin-induced MLC phosphorylation;
## between thrombin-alone
and inhibitor-thrombin groups at 60 min; ### between
inhibitor-alone groups at 2 and 60 min.
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Fig. 3.
Effect of deltamethrin on thrombin-induced EC MLC phosphorylation.
Confluent BPAECs were pretreated in M199H with vehicle (0.1% ethanol)
or 100 nM deltamethrin for 1 h and then stimulated with 100 nM thrombin
for 2, 30, and 60 min. MLC phosphorylation was determined as described
in METHODS and is expressed as a
percentage of maximum response. Data are means ± SE from 3 independent experiments. Significant difference
(P < 0.05): * compared with
maximal level of MLC phosphorylation;
# between control and
deltamethrin-pretreated groups. Similar to cyclosporin, deltamethrin
did not affect either basal or maximal MLC phosphorylation but
significantly attenuated thrombin-induced MLC dephosphorylation.
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Effect of PPase 2B inhibition on thrombin-induced EC
permeability. Verin et al. (51) have previously shown
that PPase 1 is directly involved in EC contractile responses and
regulation of EC barrier function. To examine whether PPase 2B is
similarly involved, we next determined the effect of the specific PPase 2B inhibitor deltamethrin on basal and thrombin-induced permeabilities of EC monolayers. Treatment of EC monolayers with deltamethrin for 1 h
(Fig. 4) did not affect the basal level of
EC permeability but significantly enhanced (~1.5-fold) the
thrombin-induced increases in EB-BSA flux across EC monolayers.
Although speculative, these results suggest that thrombin-inducible
type 2B PPase activity may be involved in the recovery of the EC
protective barrier after initial disruption by thrombin. The
concentration of deltamethrin required was identical for both the
enhancement of thrombin-induced EC permeability and the partial
prevention of thrombin-induced MLC dephosphorylation.

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Fig. 4.
Effect of PPase 2B inhibition on thrombin-induced EC monolayer
permeability. BPAEC monolayers (n = 11) were pretreated at time 0 with
either vehicle (0.1% ethanol) or deltamethrin (delta; 100 nM). At 60 min, cells were challenged with either vehicle (M199H alone) or 100 nM
thrombin, and permeability was determined from 130 to 180 min as
described in METHODS. * and
# Significantly different compared with control and
thrombin alone, respectively, P < 0.04. Inhibition of PPase 2B by deltamethrin did not
significantly alter basal level of EC permeability but significantly
enhanced thrombin-induced albumin clearance.
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Subcellular localization of PPase 2B in
endothelium. To further elucidate PPase 2B involvement
in the regulation of EC contractile processes, we studied the
distribution of PPases 1, 2A, and 2B in myosin-enriched and
myosin-depleted fractions [prepared as previously described by
Verin et al. (51); Fig. 5] and in
cytoskeletal fractions. For preparation of the cytoskeletal fractions,
cytosolic proteins were removed by treatment of BPAEC monolayers with
1% Nonidet P-40-150 mM NaCl extraction buffer (Fig. 5). Dishes
were washed two times with the same buffer, and the insoluble proteins remaining on dishes were solubilized in SDS sample buffer. This cell
fraction, which is enriched in actin, also includes significant amounts
of myosin and has been previously characterized as cytoskeletal (32).
Western immunoblotting analysis with specific anti-PPase antibodies
revealed that PPase 2B was present in both the cytoskeletal and
myosin-depleted cytosolic fractions but not in the myosin-enriched fraction. In contrast, PPase 1 was present in both the myosin-enriched and myosin-depleted cytosolic fractions but was absent in the cytoskeletal fraction (Fig. 6). These data
(Fig. 6A) and the previous report by
Verin et al. (51) indicate that immunoreactive PPase 2A is absent in
both contractile protein-enriched fractions but is present in EC
homogenates. Together, these results indicate that only PPase 2B, but
not type 1 or 2A, is tightly associated with the actin-enriched
cytoskeleton.

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Fig. 5.
Schema for preparation of EC fractions showing algorithm employed to
identify myosin-enriched, myosin-depleted, and cytoskeletal BPAEC
fractions. For preparation of a cytoskeletal fraction enriched in
actin, BPAECs were rinsed with PBS, and cytosolic proteins were
extracted with 50 mM Tris · HCl buffer, pH 8.0, containing 1% Nonidet P-40 (NP-40), 150 mM NaCl, and protease
inhibitors (see METHODS) for 30 min
at 4°C. Extractable proteins were discarded, and remaining
detergent-insoluble (cytoskeletal) EC fraction was solubilized in
SDS-PAGE sample buffer (31) and used for Western immunoblotting. For
preparation of myosin-enriched fractions, low-ionic-strength
homogenates obtained from confluent EC monolayers (51) were treated
with 0.1% Tween 20-0.6 M NaCl for 1 h at 4°C and centrifuged,
and resulting myosin-containing supernatants were further diluted
(10-fold) with low-ionic-strength buffer (50 mM
Tris · HCl, pH 7.0, 0.1 mM EDTA, 28 mM
-mercaptoethanol, 0.5 mM phenylmethylsulfonyl fluoride, and 2 mM
benzamidine). Precipitated myosin and associated protein fractions were
collected by centrifugation and subjected to PPase activity
determination or Western immunoblotting analysis.
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Fig. 6.
Distribution of PPases in EC fractions. PPase content in cytoskeletal
fraction (A) and myosin-enriched
(M+) and myosin-depleted (M ) EC fractions
(B) was analyzed by Western
immunoblotting. A: cytoskeletal
proteins from 100-mm dishes prepared as described in Fig. 5 were
solubilized in 3 ml of SDS sample buffer (31), boiled for 5 min,
clarified by centrifugation, and subjected to 9% SDS-PAGE followed by
Western immunoblotting. Anti-PPase 1, 2A, and 2B antibodies were used
for PPase identification. Fifty nanograms of each PPase were used as
positive controls. Amount of protein extracts loaded onto gel (~30
µl) was normalized by quantitation of Coomassie-stained gels in
preliminary experiments. B: M+ pellet
and M supernatant fractions obtained as described in Fig. 5 were
subjected to 9% SDS-PAGE. Proteins were visualized by staining gel
with Coomassie R-250 (left) or
electroblotted from gel onto a nitrocellulose filter
(right), and PPases were detected by
incubation of nitrocellulose filters with specific anti-PPase 2B
antibodies (top) or anti-PPase 1 antibodies (anti-CS1 antibodies;
bottom) followed by enhanced
chemiluminescence (ECL) detection. Equivalent amounts (~25 µl) of
initial homogenate were loaded onto gel for each fraction. Nos. on
left, positions of protein
molecular-mass markers. These data and previous experiments (50)
clearly indicate that PPases 1 and 2B but not PPase 2A are tightly
associated with EC fractions enriched in contractile proteins.
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Effect of thrombin on subcellular distribution of
PPase 2B and MLC in ECs. Because thrombin caused
stress-fiber formation, reflecting increased interaction between actin
and myosin, we next sought to examine thrombin effects on the
subcellular localization of PPase 2B (colocalized with actin) and MLC
(a part of myosin). Examination by confocal microscopy of
thrombin-challenged BPAEC monolayers (Fig. 7) revealed
that, compared with control cells, thrombin produced an increased
colocalization (yellow) of MLC (green) and PPase 2B (red), with maximal
costaining at 2 min. Although PPase 2B staining was constant, there was
a time-dependent decline in MLC staining, likely due to a decrease in
MLC antibody accessibility due to increased actin polymerization and
actin-myosin interactions. However, even at 10 min, a significant
increase of costaining between PPase 2B and MLC was observed compared
with control ECs (Fig. 7, A and
C), suggesting an increase in the
interaction between PPase 2B and MLC after thrombin stimulation.

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Fig. 7.
Effect of thrombin on intracellular localization of PPase 2B and MLC.
BPAECs grown to confluence on glass coverslips were exposed to vehicle
(A) or bovine thrombin (100 nM) for
2 (B) or 10 min
(C). Cells were fixed with
paraformaldehyde and permeabilized with 0.2% Triton as described in
METHODS. Cells were incubated with
both rabbit anti-smooth MLC antibody (1:50 in Tris-NaCl-1% albumin
buffer) and mouse anti-calcineurin antibody (1:50) overnight at
4°C. Washed cells were then secondarily stained with donkey
anti-rabbit IgG antibody (fluorescein conjugated; 1:50) and donkey
anti-mouse IgG antibody (lissamine rhodamine conjugated; 1:50).
Immunofluorescence was recorded by confocal microscopy. Cells show an
increase in costaining (yellow) of MLC (green) with PPase2B (red), with
maximal costaining at 2 min.
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PPase activity in BPAEC homogenates and cytoskeletal
fractions. A previous study by Verin et al. (51) has
characterized the involvement of a type 1 PPase in the regulation of EC
contractility and barrier function. The data described in
Effect of thrombin on subcellular distribution of
PPase 2B and MLC in ECs strongly implicate the additional involvement of PPase 2B in the regulation of
EC contractility after thrombin stimulation. To further evaluate the
myosin-specific PPase involved in agonist-inducible EC contractile responses, we studied the effect of thrombin on the PPase activity present in the homogenates and specific EC fractions.
For quantification of PPase 1 and 2A
activities in EC, 32P-labeled
phosphorylase A, a well-recognized, selective, and specific substrate
for PPases 1 and 2A in mammalian tissues, was utilized (25). To
separate PPase 1 and 2A activities of EC homogenates, we used 2 nM okadaic acid in the assay buffer, which in the diluted extracts
(<0.1 U/ml in the assay) inhibits PPase 2A completely (IC50 ~ 0.1-0.3 nM) but has
no effect on PPase 1 (IC50 = 51 nM) (8, 24). Table 1 shows that stimulation
of BPAEC monolayers with 100 nM thrombin for 10 min has no significant
effect on either PPase 1 or 2A activity in cell homogenates. Increases
in either the thrombin concentration (1 µM) or the duration of
stimulation (up to 30 min) also failed to alter EC PPase 1 and 2A
activities (data not shown). Interestingly, thrombin stimulation
actually decreased PPase 1 activity against phosphorylase A (by 30%)
in the myosin-enriched fraction (Table 1). Western immunoblotting of
control and agonist-stimulated BPAEC fractions shows that this observation can be explained by the partial release of the PPase 1 catalytic subunit from the actomyosin complex, with redistribution from
the myosin-enriched to the myosin-depleted (cytosolic) fraction (Fig.
8).

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Fig. 8.
Effect of thrombin on PPase 1 fractional distribution. M
supernatant fractions and M+ pellet fractions were prepared from BPAECs
treated with vehicle (M199H) or thrombin (100 nM) for 10 min and
subjected to 9% SDS-PAGE followed by electrotransfer to nitrocellulose
filters. PPase 1 was detected by incubation of nitrocellulose filters
with specific anti-PPase 1 antibodies followed by
125I-protein A treatment. Amount
of protein loaded onto gel in each fraction was equivalent to ~25
µl of initial homogenate. Nos. on
right, positions of protein
molecular-mass markers. Thrombin induced redistribution of PPase 1 catalytic subunit from M+ fraction to M EC fraction, a finding
that correlates well with decreased myosin-associated PPase 1 activity
noted after thrombin (Table 1).
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For quantification of
Ca2+/CaM-dependent PPase 2B
activity, we utilized 32P-histone
II-S (Sigma) phosphorylated by cAMP-dependent PKA as a PPase 2B
substrate. It was initially shown that PKA-phosphorylated histone H1
was a substrate for PPases 1 and 2A but not for PPases 2B and 2C (46).
However, it was recently demonstrated that PKA-phosphorylated histone
II-S (mixed histone fraction prepared from calf thymus by sequential
high-salt extraction, precipitation in water, acid extraction, and
reprecipitation with alcohol) had a high capacity to be
dephosphorylated by PPase 2B (11). In preliminary experiments (data not
shown), we demonstrated that under our assay conditions both purified
PPases 2A and 2B, but not PPase 1, effectively dephosphorylated PKA-phosphorylated histone II-S. However, only PPase 2B activity is
dramatically increased (>8-fold) with increasing
Ca2+ concentration (from 0.1 mM
EGTA to 0.3 mM CaCl2 in the assay buffer), whereas PPase 2A activity remains unaffected. To eliminate PPase 2A contribution in EC PPase activity against phosphohistone II-S,
we measured PPase activity in the presence of 10 nM calyculin, which we
have determined to inhibit completely both type 1 and type 2A PPases in
EC homogenates (51) but has no effect on PPases 2B and 2C (27, 45).
Although the addition of calyculin (10 nM) significantly decreased
PPase activity against histone II-S in EC homogenates by ~70% (Fig.
9A), we have assumed
that the remaining calyculin-insensitive PPase activity against histone II-S reflects PPase 2B. In contrast to the lack of induced PPase 1 activity after thrombin (Table 1), pretreatment of BPAECs with 100 nM
thrombin led to significant increases in calyculin-insensitive EC PPase
2B activity against histone II-S (Fig.
9B) in a time-dependent manner
(peaked at 20 min with a 1.7-fold increase). Consistent with these
results, BPAEC stimulation with the
Ca2+ ionophore ionomycin (10 µM)
also led to rapid activation of calyculin-insensitive Ca2+/CaM-dependent PPase 2B
activity in EC homogenates (Fig.
9A).

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Fig. 9.
Effect of agonists on histone PPase activity in EC homogenates. BPAEC
monolayers were stimulated with ionomycin (5 min, 10 µM;
A) or 100 nM thrombin for indicated
time periods (B), and PPase activity
against protein kinase (PK) A-phosphorylated histone II-S was
determined in presence and absence of 10 nM calyculin A as described in
METHODS.
A: values are means ± SE for 4 independent experiments. * Significant difference compared with
total activity in control monolayers,
P < 4 × 10 3.
# Significant difference
compared with calyculin-insensitive activity in control monolayers,
P = 4.4 × 10 3.
B: BPAECs were treated with thrombin
at time 0. PPase activity of EC
homogenates from stimulated ( ) and control ( ) cells was
sequentially monitored. Values are means ± SE for 8 independent
experiments. * Significant difference compared with control
activity (either total or calyculin-insensitive activity),
P 0.03. Thrombin stimulates
calyculin-insensitive PPase 2B activity in a time-dependent manner,
with this activation mimicked by
Ca2+ ionophore ionomycin.
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Because we detected the presence of PPase 2B but not PPase 1 or 2A in
the cytoskeleton (Fig. 10), we next
determined the effect of thrombin-induced PPase 2B activation on the
phosphorylation status of cytoskeletal proteins. We purified
cytoskeletal fractions from
32P-labeled EC monolayers and
analyzed SDS-PAGE radioactive protein profiles by autoradiography.
Figure 10 demonstrates that EC treatment with the specific PPase 2B
inhibitor deltamethrin caused a significant increase in the
phosphorylation of several cytoskeletal proteins, indicating basal
PPase 2B activity in the cytoskeleton. Thrombin treatment (100 nM, 30 min) caused the significant dephosphorylation of specific proteins in
the cytoskeleton, which was completely abolished by pretreatment with
deltamethrin. Similar results were obtained with cells treated with
ionomycin (data not shown). Taken together, these results demonstrate
the existence of thrombin-inducible PPase 2B activity that is tightly
associated with the cytoskeleton.

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Fig. 10.
Effect of thrombin on cytoskeletal protein phosphorylation. Shown is
autoradiogram of phosphorylated cytoskeletal proteins obtained from
thrombin-treated ECs labeled with
32P. BPAEC monolayers were loaded
with
[32P]orthophosphate
(0.4 mCi/ml) in phosphate-free DMEM in presence of 1% serum for 4 h
and then preincubated with vehicle (0.1% ethanol) or deltamethrin (1 µM) for 1 h after thrombin treatment (100 nM, 30 min). +,
Presence; , absence. Cytoskeletal fractions were prepared as
described in METHODS, subjected to
SDS-PAGE (31), and stained for proteins with Coomassie blue R-250.
Stained gels were dried and subjected to autoradiography with ECL
Hyperfilm. Nos. on left, positions of
molecular-mass standards. Arrows, major phosphorylated bands. Thrombin
caused a significant decrease in cytoskeletal protein phosphorylation,
which was completely attenuated by deltamethrin.
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Effect of thrombin on PPase 2B phosphorylation
status. Our data indicate that both the
Ca2+ ionophore ionomycin
and thrombin are able to activate
Ca2+/CaM-dependent PPase 2B in the
endothelium (Fig. 9). However, the time course of thrombin-induced
PPase 2B activation, which peaks at 20 min (Fig.
9B), correlates poorly with
thrombin-induced Ca2+ influx,
which peaks at <1 min and rapidly declines (18). To explore
additional factors that may modulate PPase 2B activation in
thrombin-stimulated endothelium, we examined the status of PPase 2B
phosphorylation after thrombin stimulation in denaturing immunoprecipitates from
32P-labeled BPAEC monolayers.
Figure 11 demonstrates that thrombin initially produces a rapid PPase 2B dephosphorylation. Although we were
not able to detect a rapid transient increase in PPase 2B activity in
EC homogenates after thrombin, we believe that the thrombin-induced
decrease in PPase 2B phosphorylation may result from
Ca2+-dependent PPase 2B
autodephosphorylation. The subsequent time-dependent thrombin-induced
increase in the phosphorylation status of PPase 2B correlated well with
the increase in PPase 2B catalytic activity noted in EC homogenates and
depicted in Fig. 9. These data suggest that thrombin initiates a
sequence of events that lead to time-dependent PPase 2B activation in
endothelium, possibly via phosphorylation of the enzyme. Although
information is limited about the role of PPase 2B phosphorylation in
the regulation of enzymatic activity, the activation of PPase 2B via
phosphorylation of a PPase 2B catalytic subunit has been reported (41).

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Fig. 11.
Effect of thrombin on PPase 2B catalytic subunit phosphorylation. BPAEC
monolayers from 60-mm dishes were preloaded with
[32P]orthophosphate
(0.5 mCi/ml in phosphate-free DMEM) for 2.5 h at 37°C, followed by
stimulation with either vehicle alone or thrombin (100 nM, various
times). Stimuli were then removed; monolayers were rinsed 2 times with
2 ml of medium, further rinsed with 2 ml of PBS, and scraped into 100 µl of SDS-denaturing stop solution; and cell extracts were used for
immunoprecipitation under denaturing conditions with PPase 2B-specific
antibodies as described in METHODS.
Immunocomplexes were then subjected to SDS electrophoresis (31). After
electrophoresis, proteins were transferred to nitrocellulose membranes,
and 32P signals were detected by
autoradiography at 70°C. To identify PPase 2B position,
membranes were subsequently stained with specific PPase 2B antibodies.
Relative intensities of
32P-labeled PPase 2B were
quantified by scanning densitometry.
A: Western immunoblot of purified
(Pur.) PPase 2B catalytic subunit (50 ng) and PPase 2B
immunoprecipitated from BPAECs (EC I/P). Nos. on
left, positions of molecular-mass
markers. Arrows, positions of PPase 2B and IgG.
B: autoradiogram (Autorad) of
phosphorylated PPase 2B from thrombin-treated ECs loaded with
32P
(top), an immunoblot of the same
membrane stained with anti-PPase 2B antibodies
(middle), and quantitation of level
of PPase 2B phosphorylation (mean ± SE) in 3 independent
experiments (bottom). 2',
10', and 60', 2, 10, and 60 min, respectively. Thrombin
stimulation results in a time-dependent increase in PPase 2B catalytic
subunit phosphorylation in intact BPAECs, with maximal 260% increase
at 10 min as detected by densitometric scanning.
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DISCUSSION |
There is now substantial evidence implicating MLCK-catalyzed MLC
phosphorylation as an essential element of agonist-stimulated EC
barrier dysfunction. EC contraction and the resultant intercellular gap
formation and barrier dysfunction are associated with redistribution of
microfilament proteins and are dependent on the level of MLC phosphorylation (43, 44, 52). Recently, it was shown that EC exposure
to thrombin results in isometric tension development, which correlates
with MLC phosphorylation, and is accompanied by association of myosin
with the detergent-insoluble cytoskeleton (21, 36). Consistent with
these observations, a recent study by Garcia et al. (16) in intact ECs
demonstrated that MLCK inhibition (with either ML-7 or KT-5926)
attenuates thrombin-induced MLC phosphorylation and barrier
dysfunction. Less clear, however, is the extent and type of involvement
of Ser/Thr PPase activities in the regulation of thrombin-mediated EC
MLC phosphorylation-dephosphorylation. In nonmuscle cell systems, the
addition of specific PPase inhibitors leads to changes such as actin
aggregation in the cytoskeleton (30), which implicates the involvement
of PPases in the regulation of nonmuscle contraction. In smooth muscle
tissues, PPase-mediated MLC dephosphorylation deactivates the
actomyosin ATPase, causing relaxation (see Refs. 10, 48 for reviews).
For example, the addition of a PPase-enriched protein fraction derived
from aortic smooth muscle cells (SMCs) was shown to enhance relaxation
in skinned fibers (42) and to decrease isometric force, shortening velocity, and MLC phosphorylation (2, 3). Four different PPases were
subsequently identified in gizzard smooth muscle extracts; however,
only two PPases were able to dephosphorylate intact myosin (39). A
myosin-associated PPase in skeletal, cardiac, and smooth muscles was
subsequently identified as a type 1 enzyme that was able to effectively
dephosphorylate native myosin, whereas PPase 2A was effective only
toward isolated MLC but was ineffective in dephosphorylation of native
myosin (1, 7). Recently, it was shown that several nonmuscle cells,
similar to SMCs, contain the 130-kDa "targeting subunit" of
myosin-associated PPase 1 (37). Microinjection of PPase 1 but not of
PPase 2A into mammalian fibroblasts modulated MLC phosphorylation and
induced microfilament reorganization (12), which suggests that, similar
to SMCs, PPase 1 is involved in the regulation of nonmuscle cell
contractility. Using primarily inhibitory analysis, Verin et al. (51)
previously reported that, similar to SMCs, the myosin PPase in ECs
involved in physiologically relevant MLC dephosphorylation is a type 1 enzyme. Consistent with our data, Shinoki et al. (47) demonstrated that
PPase 1 plays a predominant role in sustaining the normal EC
cytoskeletal structure.
Although these studies suggest the primacy of a type 1 PPase in MLC
dephosphorylation in mammalian muscle and nonmuscle systems, Ca2+-dependent PPase 2B purified
from scallop smooth muscle was shown to effectively dephosphorylate MLC
bound in scallop opaque and aortic smooth muscle myosin (26) and to be
involved in the maintenance of catch tension by MLC dephosphorylation
in molluscan muscles (6, 26). PPase 2B has been shown to participate in
sperm motility and to be tightly associated with mammalian nonmuscle cytoskeleton (13, 38, 49). These reports raise the possibility that
PPase 2B may also be involved in the regulation of contractility in
specific cell types. The present study focused on the role of
Ca2+/CaM-dependent PPase 2B in
agonist-stimulated EC contractile responses. Our present results
clearly demonstrate that thrombin (100 nM) causes significant
(>2-fold) and rapid (2-min) increases in MLC phosphorylation, which
gradually declined, with dephosphorylation nearly complete after 1 h
(Fig. 1). Garcia et al. (16) previously showed that higher doses of
thrombin (1 µM) caused a three- to fourfold increase in MLC
phosphorylation (~1.2 mol phosphate/mol MLC) above basal constitutive
levels, with maximal effect at 1 min, followed by MLC
dephosphorylation, which was nearly complete by 30 min. Taken together,
these data demonstrate that thrombin has dose-dependent effects on EC
MLC phosphorylation-dephosphorylation status and implicates the
involvement of Ser/Thr PPases in EC MLC dephosphorylation events after
thrombin stimulation. Although we clearly determined a type 1 PPase is
involved in thrombin-mediated MLC dephosphorylation, calyculin A, a
potent inhibitor of both type 1 and type 2A PPases (24, 27), failed to
prevent MLC dephosphorylation after thrombin (Fig.
2A), suggesting that
calyculin-insensitive PPase activities may be involved in MLC
dephosphorylation after thrombin.
The two major types of calyculin-insensitive PPases that exist in cells
are highly dependent on divalent cation availability: Ca2+/CaM-dependent PPase 2B and
Mg2+-dependent PPase 2C (44).
Although there is no evidence that agonists such as thrombin affect the
intracellular Mg2+ concentration
in the endothelium, Garcia et al. (18) and others (22) have previously
shown that thrombin-mediated EC activation caused rapid (<15-s)
transient increases in intracellular
Ca2+ concentration. Thus it is
logical to assume that PPase 2B rather than PPase 2C would be more
likely involved in thrombin-mediated EC contractile responses. The
specific PPase 2B inhibitors cyclosporin (Fig.
2B) and deltamethrin (Fig. 3), at
concentrations sufficient for complete PPase 2B inhibition (11, 14),
did not alter the level of either basal or maximal thrombin-induced MLC
phosphorylation but significantly attenuated the predicted MLC
dephosphorylation after thrombin. Furthermore, inhibition of
thrombin-induced MLC dephosphorylation by deltamethrin was accompanied
by a subsequent increase in thrombin-induced EC permeability (Fig. 4).
Although not definitive, the same concentration of PPase 2B inhibitor
was equally effective for both events, suggesting that the effects on
MLC phosphorylation and permeability may be linked. Several studies
have previously implicated the involvement of PPase 2B in EC activation
responses. For example, PPase 2B is involved in endothelin-1 gene
regulation in human endothelium (35), and prolonged incubation of human
ECs with cyclosporin A (4 days, 0.3-7 µM) results in significant
cell detachment (4). To our knowledge, our data represent the first
demonstration that PPase 2B may have an important regulatory role in EC
contractile and barrier responses.
To further elucidate PPase 2B involvement in the regulation of EC
contractile processes, we studied the distribution of this PPase in
different EC fractions. Our results indicate that immunoreactive PPase
2B is present in a cytoskeletal actin-enriched fraction (which includes
significant amounts of myosin) and in the myosin-depleted cytosolic
fraction. In contrast, PPase 1 is present in the myosin-enriched and
myosin-depleted fractions but is entirely absent in the
detergent-insoluble cytoskeleton. Taken together with previous
observations by Verin et al. (51), these results suggest that PPase 2B
but not PPase 1 or 2A is tightly associated with the EC cytoskeleton.
In agreement with our data, highly active PPase 2B was recently
purified from the detergent-insoluble adrenal cell cytoskeleton (38),
and neuronal cell treatment with cytochalasin D resulted in
dissociation of PPase 2B from disrupted actin filaments (13). These
observations suggest that the association of PPase 2B with the
cytoskeleton depends on intact actin filaments (13) rather than on
filamentous myosin, although we cannot exclude the possibility that
PPase 2B can be specifically associated with a specific myosin heavy chain II isoform (A or B) that is present in nonmuscle cells (28).
It has been recently reported that PPase 1 is also associated with the
detergent-insoluble cytoskeletal fraction from cerebral cortex (9). We
did not find immunoreactive PPase 1 associated with the EC
cytoskeleton; however, previously published observations by Verin et
al. (51) indicated that ~10% of total phosphorylase PPase activity
(combined PPase 1 and 2A activity) resides in the cytoskeletal
fraction. Western blotting analysis of EC fractions with anti-PPase 1 antibodies produced against the type 1 catalytic subunit-
(CS1
)
(Fig. 7A) and anti-CS1
isoform
antibodies (data not shown) did not detect any CS1 protein band in the
cytoskeletal fraction. Therefore, these data suggest that the PPase 1 activity associated with the EC cytoskeleton may be attributable to a
CS1 isoform that is immunologically distinct from the CS1
and -
isoforms.
Our data indicate that PPase 1 and 2A activities in EC homogenates are
not affected by thrombin stimulation (Table 1). Nevertheless, thrombin
caused a partial translocation of the EC CS1 from the myosin-enriched
to the myosin-depleted fraction (Fig. 8), which was accompanied by
decreasing myosin-associated PPase 1 activity (Table 1). These data are
consistent with the previously published observations (45) that
thrombin inhibits calyculin-sensitive MLC dephosphorylation in the
endothelium. The mechanism responsible for thrombin-induced CS1
translocation is unclear; however, dissociation of CS1 from its
targeting subunit was proposed as a mechanism of regulation of the
glycogen-associated form of PPase 1 (46). In SMCs, arachidonic acid, an
intermediate product of Gs protein activation, caused dissociation of CS1 from the myosin-associated PPase
1 holoenzyme, which may account for the G protein-mediated sensitization of the SMC contractile apparatus to
Ca2+ (23, 29). Garcia et al. (17)
previously showed that G protein-mediated EC activation by thrombin
results in Ca2+ mobilization;
phospholipase A2, C, and D
activation; and release of arachidonic acid (17). Although speculative,
it is possible that, similar to SMCs, arachidonate generated by
thrombin-stimulated phospholipase
A2 and D activity can cause CS1
translocation to the cytosol and decrease myosin-specific PPase 1 activity. Nevertheless, it is clear that the observed decrease in
myosin-associated PPase 1 activity cannot explain EC MLC
dephosphorylation after thrombin stimulation.
Our results demonstrated that thrombin caused dephosphorylation of
several proteins in the cytoskeletal fractions (Fig. 10) and that this
reaction was totally blocked by the specific PPase 2B inhibitor
deltamethrin. These data are consistent with the existence of a
thrombin-inducible PPase 2B, which is tightly bound to the contractile
apparatus. As evidenced by immunofluorescence experiments (Fig. 7),
thrombin also increased colocalization of PPase 2B and MLC, thereby
enhancing MLC substrate accessibility to PPase 2B. Finally, thrombin
significantly increased calyculin-insensitive PPase 2B activity in EC
homogenates in a time-dependent manner. This activation was evident
after 10 min of stimulation and was maximal after 20 min of thrombin
treatment (Fig. 9B), a time frame strongly correlated with MLC dephosphorylation induced by thrombin (Fig. 2). EC monolayers exposed to the
Ca2+ ionophore ionomycin (5 µM
for 15 min) also significantly increased PPase 2B activity in EC
homogenates (Fig. 9A), consistent
with the involvement of Ca2+
influx in thrombin-induced EC PPase 2B activation. Of potential concern
is the discrepancy in the time course of thrombin-induced PPase 2B
activation (Fig. 9) with the time course of the previously reported
increase in thrombin-mediated intracellular
Ca2+ concentration (18). However,
our data indicate a strong correlation between PPase 2B catalytic
subunit phosphorylation induced by thrombin and PPase 2B activity (Fig.
11). These results indicate that thrombin induces a sequence of
biochemical events, including Ca2+
mobilization and PK(s) activation, which precedes and may initiate Ca2+/CaM-dependent PPase 2B
activation in the endothelium. Taken together with the role of PPase 2B
in MLC dephosphorylation in intact EC monolayers, these experimental
data strongly implicate the involvement of inducible
cytoskeletal-associated PPase 2B activity in the regulation of
thrombin-mediated EC contractility and barrier function.
 |
ACKNOWLEDGEMENTS |
We gratefully acknowledge Lakshmi Natarajan and Lucy Robles Rivera
for superb technical assistance and Rebecca Snyder for expert
secretarial assistance. We also thank Drs. Anna DePaoli-Roach, Susan
Gunst, and James Stull for the generous supply of specific antibodies.
 |
FOOTNOTES |
This work was supported by National Heart, Lung, and Blood Institute
Grants HL-50533 and HL-57402; the American Lung Association of Indiana;
the American Heart Association National and Indiana Affiliates; and the
Department of Veterans Affairs Medical Research Service.
Address for reprint requests and present address of A. D. Verin: The
Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Circle,
Baltimore, MD 21224.
Received 16 October 1996; accepted in final form 1 July 1998.
 |
REFERENCES |
1.
Alessi, D.,
L. K. MacDougall,
M. M. Sola,
M. Ikebe,
and
P. Cohen.
The control of protein phosphatase-1 by targeting subunits. The major myosin phosphatase in avian smooth muscle is a novel form of protein phosphatase-1.
Eur. J. Biochem.
210:
1023-1035,
1992[Abstract].
2.
Bialojan, C.,
L. Merkel,
J. C. Ruegg,
D. Gilford,
and
J. DiSalvo.
Prolonged relaxation of detergent-skinned smooth muscle involves decreased endogenous phosphatase activity.
Proc. Soc. Exp. Biol. Med.
178:
648-652,
1985[Abstract].
3.
Bialojan, C.,
J. C. Ruegg,
and
J. DiSalvo.
Phosphatase-mediated modulation of actin-myosin interaction in bovine aortic actomyosin and skinned porcine carotid artery.
Proc. Soc. Exp. Biol. Med.
178:
36-45,
1985[Abstract].
4.
Bombeli, T.,
M. Muller,
P. W. Straub,
and
A. Haeberli.
Cyclosporine-induced detachment of vascular endothelial cells initiates the intrinsic coagulation system in plasma and whole blood.
J. Lab. Clin. Med.
127:
621-634,
1996[Medline].
5.
Bradford, M. M.
A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem.
72:
248-254,
1976[Medline].
6.
Castellani, L.,
and
C. Cohen.
A calcineurin-like phosphatase is required for catch contraction.
FEBS Lett.
309:
321-326,
1992[Medline].
7.
Chisholm, A. A. K.,
and
P. Cohen.
The myosin-bound form of protein phosphatase 1 (PP-1m) is the enzyme that dephosphorylates native myosin in skeletal and cardiac muscles.
Biochim. Biophys. Acta
971:
163-169,
1988[Medline].
8.
Cohen, P.,
S. Klumpp,
and
D. L. Schelling.
An improved procedure for identifying and quantitating protein phosphatases in mammalian tissues.
FEBS Lett.
250:
596-600,
1989[Medline].
9.
De Freitas, M. S.,
A. G. de Mattos,
M. M. Camargo,
C. M. D. Wannmacher,
and
R. Pessoa-Pureur.
Cytoskeletal-associated protein kinase and phosphatase activities from cerebral cortex of young rats.
Neurochem. Res.
20:
951-956,
1995[Medline].
10.
De Lanerolle, P.,
and
R. J. Paul.
Myosin phosphorylation/dephosphorylation and regulation of airway smooth muscle contractility.
Am. J. Physiol.
261 (Lung Cell. Mol. Physiol. 5):
L1-L14,
1991[Abstract/Free Full Text].
11.
Enan, E.,
and
F. Matsumura.
Specific inhibition of calcineurin by type II synthetic pyrethroid insecticides.
Biochem. Pharmacol.
43:
1777-1784,
1992[Medline].
12.
Fernandez, A.,
D. L. Brautigan,
M. Mumby,
and
N. J. C. Lamb.
Protein phosphatase type-1, not type-2A, modulates actin microfilament integrity and myosin light chain phosphorylation in living nonmuscle cells.
J. Cell Biol.
111:
103-112,
1990[Abstract].
13.
Ferreira, A.,
R. Kincaid,
and
K. S. Kosik.
Calcineurin is associated with the cytoskeleton of cultured neurons and has a role in the acquisition of polarity.
Mol. Biol. Cell
4:
1225-1238,
1993[Abstract].
14.
Fruman, D. A.,
C. B. Klee,
B. E. Bierer,
and
S. J. Burakoff.
Calcineurin phosphatase activity in T lymphocytes is inhibited by FK 506 and cyclosporin A.
Proc. Natl. Acad. Sci. USA
89:
3686-3690,
1992[Abstract].
15.
Gallagher, P. J.,
J. G. N. Garcia,
and
B. P. Herring.
Expression of a novel myosin light chain kinase in embryonic tissues and cultured cells.
J. Biol. Chem.
270:
29090-29095,
1995[Abstract/Free Full Text].
16.
Garcia, J. G. N.,
H. W. Davis,
and
C. E. Patterson.
Regulation of endothelial cell gap formation and barrier dysfunction: role of myosin light chain phosphorylation.
J. Cell. Physiol.
163:
510-522,
1995[Medline].
17.
Garcia, J. G. N.,
J. W. Fenton II,
and
V. Natarajan.
Thrombin stimulation of human endothelial cell phospholipase D activity. Regulation by phospholipase C, protein kinase C, and cyclic adenosine 3'5'-monophosphate.
Blood
79:
2056-2067,
1992[Abstract].
18.
Garcia, J. G. N.,
C. E. Patterson,
C. Bahler,
J. Aschner,
C. M. Hart,
and
D. English.
Thrombin receptor activating peptides induce Ca2+ mobilization, barrier dysfunction, prostaglandin synthesis, and platelet-derived growth factor mRNA expression in cultured endothelium.
J. Cell. Physiol.
156:
541-549,
1993[Medline].
19.
Garcia, J. G. N.,
and
K. L. Schaphorst.
Regulation of endothelial cell gap formation and paracellular permeability.
J. Investig. Med.
43:
117-126,
1995[Medline].
21.
Goeckeler, Z. M.,
and
R. B. Wysolmerski.
Myosin light chain kinase-regulated endothelial cell contraction: the relationship between isometric tension, actin polymerization, and myosin phosphorylation.
J. Cell Biol.
130:
1-15,
1995[Abstract].
22.
Goligorsky, M. S.,
D. N. Menton,
A. Laszlo,
and
H. Lum.
Nature of thrombin-induced sustained increase in cytosolic calcium concentration in cultured endothelial cells.
J. Biol. Chem.
264:
16771-16775,
1989[Abstract/Free Full Text].
23.
Gong, M. C.,
A. Fuglsang,
D. Alessi,
S. Kobayashi,
P. Cohen,
A. V. Somlyo,
and
A. P. Somlyo.
Arachidonic acid inhibits myosin light chain phosphatase and sensitizes smooth muscle to calcium.
J. Biol. Chem.
267:
21492-21498,
1992[Abstract/Free Full Text].
24.
Honkanan, R. E.,
B. A. Codispoti,
K. Tse,
and
A. L. Boynton.
Characterization of natural toxins with inhibitory activity against serine/threonine protein phosphatases.
Toxicon
32:
339-350,
1994[Medline].
25.
Ingebritsen, T. S.,
A. A. Stewart,
and
P. Cohen.
The protein phosphatases involved in cellular regulation. 6. Measurement of type-1 and type-2 protein phosphatases in extracts of mammalian tissues; an assessment of their physiological roles.
Eur. J. Biochem.
132:
297-307,
1983[Abstract].
26.
Inoue, K.,
H. Sohma,
and
F. Morita.
Ca2+-dependent protein phosphatase which dephosphorylates regulatory light chain-a in scallop smooth muscle myosin.
J. Biochem. (Tokyo)
107:
872-878,
1990[Abstract].
27.
Ishihara, H.,
B. L. Martin,
D. L. Brautigan,
H. Karaki,
H. Ozaki,
Y. Kato,
N. Fusetani,
S. Watabe,
K. Hashimoto,
D. Vemura,
and
D. J. Hartshorne.
Calyculin A and okadaic acid: inhibitors of protein phosphatase activity.
Biochem. Biophys. Res. Commun.
106:
1126-1133,
1989.
28.
Kawamoto, S.,
and
R. S. Adelstein.
Chicken nonmuscle myosin heavy chains: differential expression of two mRNAs and evidence for two different polypeptides.
J. Cell Biol.
112:
915-924,
1991[Abstract].
29.
Kitazawa, T.,
M. Masuo,
and
A. P. Somlyo.
G protein-mediated inhibition of myosin light-chain phosphatase in vascular smooth muscle.
Proc. Natl. Acad. Sci. USA
88:
9307-9310,
1991[Abstract].
30.
Kreinbühl, P.,
H. Keller,
and
V. Niggli.
Protein phosphatase inhibitors okadaic acid and calyculin A alter cell shape and F-actin distribution and inhibit stimulus-dependent increases in cytoskeletal actin of human neutrophils.
Blood
80:
2911-2919,
1992[Abstract].
31.
Laemmli, U. K.
Cleavage of structural proteins during the assembly of head of bacteriophage.
Nature
227:
680-685,
1970[Medline].
32.
Lee, W.-C.,
J.-S. Yu,
S.-D. Yang,
and
Y.-K. Lai.
Reversible hyperphosphorylation and reorganization of vimentin intermediate filaments by okadaic acid in 9L rat brain tumor cells.
J. Cell. Biochem.
49:
378-393,
1992[Medline].
34.
Lum, H.,
and
A. B. Malik.
Regulation of vascular endothelial barrier function.
Am. J. Physiol.
267 (Lung Cell. Mol. Physiol. 11):
L223-L241,
1994[Abstract/Free Full Text].
35.
Marsen, T. A., M. S. Simonson, and M. J. Dunn. Thrombin-mediated ET-1 gene regulation involves CaM
kinases and calcineurin in human endothelial cells. J. Cardiovasc. Pharmacol. 6, Suppl. 3: S1-S4, 1995.
36.
Moy, A. B.,
J. Van Engelenhoven,
J. Bodmer,
J. Kamath,
C. Keese,
I. Giaever,
S. Shasby,
and
D. M. Shasby.
Histamine and thrombin modulate endothelial focal adhesion through centripetal and centrifugal forces.
J. Clin. Invest.
97:
1020-1027,
1996[Abstract/Free Full Text].
37.
Okubo, S.,
M. Ito,
Y. Takashiba,
K. Ichikawa,
M. Miyahara,
H. Shimizu,
T. Konishi,
H. Shima,
M. Nagao,
D. J. Hartshorne,
and
T. Nakano.
A regulatory subunit of smooth muscle myosin bound phosphatase.
Biochem. Biophys. Res. Commun.
200:
429-434,
1994[Medline].
38.
Papadopoulos, V.,
A. S. Brown,
and
P. Hall.
Isolation and characterization of calcineurin from adrenal cell cytoskeleton: identification of substrates for Ca2+-calmodulin-dependent phosphatase activity.
Mol. Cell. Endocrinol.
63:
23-38,
1989[Medline].
39.
Pato, M. D.,
and
E. Kerk.
Purification and characterization of a smooth muscle myosin phosphatase from turkey gizzards.
J. Biol. Chem.
260:
12359-12366,
1985[Abstract/Free Full Text].
40.
Patterson, C. E.,
R. A. Rhoades,
and
J. G. N. Garcia.
Evans blue-albumin binding as a new marker of transendothelial macromolecule permeability.
J. Appl. Physiol.
72:
865-873,
1992[Abstract/Free Full Text].
41.
Pereira, M. M. C.,
D. K. Shori,
R. L. Dormer,
and
M. A. McPherson.
Studies on phosphorylation of calcineurin.
Biochem. Soc. Trans.
18:
447,
1990[Medline].
42.
Ruegg, J. C.,
J. DiSalvo,
and
R. J. Paul.
Soluble relaxation factor from vascular smooth muscle: a myosin light chain phosphatase.
Biochem. Biophys. Res. Commun.
106:
1126-1133,
1982[Medline].
43.
Schnittler, H. J.,
A. Wilke,
T. Gress,
N. Suttorp,
and
D. Drenckhahn.
Role of actin and myosin in the control of paracellular permeability in pig, rat, and human vascular endothelium.
J. Physiol. (Lond.)
431:
379-401,
1990[Abstract].
44.
Shasby, D. M.,
S. S. Shasby,
J. M. Sullivan,
and
M. J. Peach.
Role of endothelial cell cytoskeleton in control of endothelial permeability.
Circ. Res.
51:
657-661,
1982[Abstract].
45.
Shasby, D. M.,
T. Stevens,
D. Ries,
A. B. Moy,
J. M. Kamath,
A. M. Kamath,
and
S. S. Shasby.
Thrombin inhibits myosin light chain dephosphorylation in endothelial cells.
Am. J. Physiol.
272 (Lung Cell. Mol. Physiol. 16):
L311-L319,
1997[Abstract/Free Full Text].
46.
Shenolikar, S.,
and
A. C. Nairn.
Protein phosphatases: recent progress.
Adv. Second Messenger Phosphoprotein Res.
23:
1-121,
1991[Medline].
47.
Shinoki, N.,
M. Sakon,
J. Kambayashi,
M. Ikeda,
E. Oijiki,
M. Okuyama,
K. Fujitani,
Y. Yano,
T. Kawasaki,
and
M. Monden.
Involvement of protein phosphatase-1 in cytoskeletal organization of cultured endothelial cells.
J. Cell. Biochem.
59:
368-375,
1995[Medline].
48.
Somlyo, A. P.,
and
A. V. Somlyo.
Signal transduction and regulation in smooth muscle.
Nature
372:
231-236,
1994[Medline].
49.
Tash, J. S.,
M. Krinks,
J. Patel,
R. L. Means,
C. B. Klee,
and
A. R. Means.
Identification, characterization, and functional correlation of calmodulin-dependent protein phosphatase in sperm.
J. Cell Biol.
106:
1625-1633,
1988[Abstract].
50.
Towbin, H.,
T. Staehelin,
and
J. Gordon.
Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications.
Proc. Natl. Acad. Sci. USA
76:
4350-4354,
1979[Abstract].
51.
Verin, A. D.,
C. E. Patterson,
M. A. Day,
and
J. G. N. Garcia.
Regulation of endothelial cell gap formation and barrier function by myosin-associated phosphatase activities.
Am. J. Physiol.
269 (Lung Cell. Mol. Physiol. 13):
L99-L108,
1995[Abstract/Free Full Text].
52.
Wysolmerski, R. B.,
and
D. Lagunoff.
Regulation of permeabilized endothelial cell retraction by myosin phosphorylation.
Am. J. Physiol.
261 (Cell Physiol. 30):
C32-C40,
1991[Abstract/Free Full Text].
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