Divisions of 1Infectious Diseases and 4Pulmonary and Critical Care Medicine, 2Departments of Medicine and Pathology, Department of Veterans Affairs Medical Center, 5Mucosal Biology Research Center, University of Maryland School of Medicine, Baltimore 21201; and 3Division of Pulmonary and Critical Care Medicine, 6Departments of Medicine, Anesthesiology, Cell Biology, and Pediatrics, Johns Hopkins University School of Medicine, Baltimore, Maryland 21224
Submitted 11 December 2002 ; accepted in final form 24 February 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
zonula adherens; angiogenesis; endothelial barrier function; vascular endothelial cadherin; catenins
In the EC, a circumferential band of actin microfilaments
(19,
58) is tethered to the EC-EC
adherens junction or the zonula adherens (ZA), an intercellular junctional
complex that modulates homophilic cell-cell adhesion
(18,
44). Vascular endothelial
(VE)-cadherin, a membrane-spanning glycoprotein with an ectodomain that
dictates homophilic adhesive specificity and a cytoplasmic domain that is
indirectly tethered to the actin cytoskeleton, is central to ZA organization
(18,
44). Although multiple
cadherins can be coexpressed and differentially distributed in EC, VE-cadherin
appears to be unique in that it is localized to the intercellular junctions
(17,
18,
44). At least three
cytoplasmic proteins collectively termed the catenins, including
-catenin,
-catenin,
-catenin, also known as plakoglobin,
and possibly p120ctn, participate in anchoring the cytoplasmic
domain of cadherins to actin microfilaments.
-,
-, And p120
catenin each directly binds to cadherin.
- And
-catenin compete
for the same binding site, whereas p120ctn associates with a more
membrane-proximal sequence.
- And
-catenin each bind to
-catenin (1,
51), which couples the
cadherin-catenin complex to the actin cytoskeleton
(40,
51). This ZA/peripheral actin
band forms a continuous belt around the apical portion of the cell, where it
is strategically localized to modulate EC-EC interactions and the paracellular
pathway (17,
19).
The state of ZA protein tyrosine phosphorylation is central to the
regulation of the ZA/actin cytoskeletal linkage and homophilic cell-cell
adhesion (33,
34,
39) and, as we
(31,
60) and others
(20,
46,
53) have shown, to
angiogenesis and the maintenance of endothelial barrier function. Although the
signal transduction pathways that regulate the state of ZA assembly are
incompletely understood, ZA proteins can be modified through tyrosine
phosphorylation. -,
-,
- And p120-catenin, and cadherins
themselves each can be phosphorylated on tyrosine residues
(1,
20,
33,
54). Multiple stimuli that
induce tyrosine phosphorylation of ZA proteins, including src and ras
transformation, mitogenic growth factors, proangiogenic agonists,
counteradhesive proteins, and cytokines, profoundly alter their organization.
Increased tyrosine phosphorylation of one or more of these proteins,
especially
-catenin, reduces cadherin ectodomain homophilic adhesion. In
certain cases, increased tyrosine phosphorylation of
-catenin can
promote disassembly of the ZA complex, and/or uncoupling of the ZA from the
actin cytoskeleton (34,
39).
Protein tyrosine phosphatases (PTPs) are thought to play a crucial role in regulating the state of ZA protein tyrosine phosphorylation and assembly. Increased expression of a number of PTPs parallels increases in cell density (13, 25, 50). As cells achieve confluence, a group of receptor PTPs that participate in strict homophilic adhesion with identical molecules expressed on the surface of neighboring cells is sequestered at the cell-cell interface (10, 15, 23, 28). Here, their catalytic activities are strategically localized and sustained in close proximity to intercellular junctions, including the ZA. Several PTPs directly associate with and/or dephosphorylate ZA proteins (6, 12, 15, 23, 55).
A number of studies support a role for PTPs in the maintenance of EC-EC
junctional integrity and endothelial barrier function. In contact-inhibited
confluent human umbilical vein EC (HUVEC), membrane-associated PTP activity is
increased 12-fold compared with subconfluent EC
(25). One or more PTPs are
prominently or almost exclusively expressed in endothelia
(9,
24,
55). In mouse development
(24) and adult
(9) studies, PTP-µ
transcription is almost exclusively localized to vascular endothelia where it
colocalizes with Flk-1, a receptor PTK that is exclusively expressed in EC. In
HUVEC, the SH2 domain-containing PTP, SHP-2, binds to
-catenin and
restrains phosphorylation of
-,
-, and p120 catenins
(55). More recently, VE-PTP,
the murine homolog of human PTP-
, was shown to associate with
VE-cadherin through its ectodomain and to regulate VE-cadherin phosphorylation
and barrier-enhancing activity in transfected Chinese hamster ovary cells,
independently of its PTP domain
(48). We have found that, for
some mediators of tyrosine phosphorylation-dependent increments in
paracellular permeability, concurrent PTP inhibition is required for a
reliable, reproducible phosphotyrosine signal
(31,
61). We also found that
coadministration of two structurally and functionally dissimilar PTP
inhibitors, vanadate and phenylarsine oxide (PAO), at levels that do not in
themselves alter barrier function, enhanced transendothelial albumin flux in
response to multiple agonists. In one study, nonselective PTP inhibition
decreased electrical resistance across bovine brain EC monolayers coincident
with tyrosine phosphorylation of proteins associated with intercellular
junctions (53). In a more
recent study, diperoxovanadate, a potent PTK activator and PTP inhibitor,
decreased transendothelial electrical resistance and increased tyrosine
phosphorylation of ZA proteins in bovine pulmonary artery EC
(27). Finally, several
mediators of increased vascular permeability, including thrombin, TNF-
,
and IL-1, reportedly modulate PTP activity
(2,
29,
32,
55).
Irrespective of the receptor or signaling pathway activated, a final common pathway for increased endothelial paracellular permeability appears to exist, in which decreased PTP catalytic function permits increased tyrosine phosphorylation of ZA proteins and decreased VE-cadherin ectodomain-mediated, homophilic adhesion. In the current study, we asked whether nonselective PTP inhibition that circumvents agonist activation of one or more EC receptors might disrupt EC-EC junctional integrity in postconfluent EC monolayers or preformed EC tubes and whether these effects were associated with tyrosine phosphorylation of one or more ZA proteins, and/or disassembly of the VE-cadherin-catenin complex.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Assay of transendothelial albumin flux. Transendothelial [14C]bovine serum albumin (BSA) flux was assayed as previously described (7, 31). Briefly, gelatin-impregnated polycarbonate filters (13-mm diameter, 0.4-µm pore size) (Nucleopore, Pleasanton, CA) mounted in polystyrene chemotactic chambers (ADAPS, Dedham, MA) were inserted into wells of 24-well plates. EC (2 x 105) were seeded in each upper compartment and were cultured for 72 h. We established the baseline barrier function of each monolayer by introducing an equivalent concentration of permeability tracer [14C]BSA (1.1 pmol, i.e., 4,8006,200 dpm/0.5 ml) to each upper compartment for 1 h at 37°C, after which 0.5 ml from the lower compartment was mixed with 4.5 ml of Optifluor Scintillation fluid (Packard Instruments, Downers Grove, IL) and counted in a liquid scintillation counter (Packard). Only those monolayers retaining >97% of the tracer were studied. The monolayers were exposed for 5 min6 h to the nonselective PTP inhibitors sodium orthovanadate (vanadate, Sigma) or PAO (Sigma) or to medium alone. PTP inhibition was also studied in the presence of the PTK inhibitor herbimycin A (1.0 µM, Sigma), which was introduced 16 h before and sustained throughout the vanadate treatment.
Assay of EC injury. To determine whether vanadateor PAO-induced barrier dysfunction could be ascribed to EC injury, a 51Cr release assay was employed as previously described (31). Briefly, EC were labeled with [51Cr]sodium chromate (Amersham, Arlington, IL), and the labeled monolayers were incubated with vanadate, PAO, or medium alone. The supernatants were centrifuged and counted. All washed monolayers were solubilized with 1% Triton X-100 (Sigma) to induce maximum release. The lysates were centrifuged and the supernatants counted for 51Cr activity. EC injury was expressed as [(51Cr supernatant)/(51Cr supernatant + 51Cr cell lysate)] x 100%.
EC tube formation assay. Each well of a six-well plate was coated with 0.2 ml of Matrigel (8 mg/ml; BD Biosciences, Bedford, MA) as previously described (60). EC were seeded at 4 x 105 cells/well into the Matrigel-coated wells and cultured for 16 h in DMEM containing 10% FBS to allow for tube formation. The EC were then incubated for an additional 6 h with vanadate (25 µM), PAO (0.25 µM), or media alone. Tubular structures were photographed with a Zeiss inverted microscope and a x32 objective.
F-actin epifluorescence microscopy and immunolocalization of
phosphotyrosine-containing proteins and ZA components. To maintain EC
monolayers under experimental conditions identical to those of our barrier
function assay, we stained monolayers directly on polycarbonate filters as
previously described (7).
Briefly, EC cultured to confluence on these filters were exposed to vanadate,
PAO, or media alone. In some experiments, monolayers were fixed, rendered
permeable, and stained with fluorescein-phalloidin (1.65 x
10-7 M; Molecular Probes, Eugene, OR). The filters and
their attached monolayers were photographed through a Zeiss Axioskop 20
microscope equipped for epifluorescence. In other experiments, EC cultured on
filters in the presence of vanadate or media alone were probed for
phosphotyrosine-containing proteins with fluorescein isothiocyanate
(FITC)-conjugated antiphosphotyrosine antibody (5 µg/ml in PBS containing
1% BSA; Upstate Biotechnology, Lake Placid, NY) as described
(61). For colocalization of
VE-cadherin with either - or
-catenin, EC were cultured to
postconfluence on fibronectin-coated glass coverslips, after which they were
incubated for increasing times with increasing concentrations of vanadate or
media alone. The EC were washed, fixed, permeabilized with 3% paraformaldehyde
plus 0.5% Triton X-100 for 2 min followed by paraformaldehyde for 20 min, and
incubated for 0.5 h with rabbit anti-human VE-cadherin IgG (ICOS, Bothell, WA)
and either murine anti-human
-catenin or
-catenin antibodies
(Transduction Laboratories, Lexington, KY). This was followed by incubation
with affinity cross-adsorbed secondary antibodies including FITC-conjugated
donkey anti-rabbit IgG (Chemicon, Temecula, CA) and Cy5 (near-infrared
emission)-conjugated donkey anti-mouse IgG (Jackson Immunochemicals, West
Grove, PA). These samples were photographed through a Nikon TE-200 microscope
with x60 plan-apo objective with a numerical aperture of 1.4. Images
were captured with a quadruple dichroic mirror and separate excitation and
emission filters for fluorescein and near-infrared fluorescence (Chroma,
Portland, ME) mounted in Orbit filter wheels (Manchester, UK), and a Roper HQ
charge-coupled device camera, all run with Openlab software (Improvision,
Lexington, MA).
Immunoblotting for EC phosphotyrosine. EC (2.3 x
105 cells/100-mm dish) cultured to confluence were exposed to
vanadate or to media alone after which they were lysed with ice-cold lysis
buffer containing 50 mM Tris · HCl (pH 7.4), 1% Nonidet P-40, 0.25%
sodium deoxycholate, 150 mM NaCl, 1 mM phenylmethylsulfonyl fluoride, 1 mg/ml
leupeptin, 1 mg/ml pepstatin A, 1 mg/ml aprotinin, 1 mM vanadate, 1 mM sodium
fluoride, 10 mM disodium pyrophosphate, 500 µM paranitrophenol, and 1 mM
PAO (all purchased from Sigma). The EC lysates were assayed for protein
concentration with a Bio-Rad DC Protein assay kit (Bio-Rad, Richmond, CA).
Samples were resolved by electrophoresis on an 816% gradient sodium
dodecyl sulfate (SDS)-polyacrylamide gel (Novex, San Diego, CA) and
transferred onto polyvinylidene difluoride (PVDF) membranes (Millipore,
Bedford, MA). The blots were probed with a biotinylated antiphosphotyrosine
monoclonal antibody (0.8 µg/ml, 4G10; Upstate Biotechnology), incubated
with horseradish peroxidase (HRP)-conjugated streptavidin (0.5 µg/ml,
Upstate Biotechnology), and developed with enhanced chemiluminescence (ECL,
Amersham). To confirm equivalent protein loading and transfer, we stripped
blots with 100 mM -mercaptoethanol, 2% SDS, and 62.5 mM Tris ·
HCl, pH 6.7, reprobed them with 0.5 µg/ml murine anti-physarum
-tubulin IgG2b (Boehringer-Mannheim, Indianapolis, IN)
(31,
61), and developed them as
described above. Blots were scanned by laser densitometry (Molecular Dynamics,
Sunnyvale, CA), and phosphotyrosine signal was normalized to
-tubulin.
Identification of phosphotyrosine-containing proteins. An
immunoprecipitation strategy was employed to identify substrates for tyrosine
phosphorylation as previously described
(31). Lysates of EC treated
with vanadate or media alone were precleared by incubation with either
anti-murine or anti-goat IgG cross-linked to agarose (Sigma) for 1 h at
4°C and then incubated overnight at 4°C with specific murine
monoclonal antibodies raised against -,
-, or p120-catenin
(Transduction Laboratories), or a goat polyclonal antibody raised against
VE-cadherin (Santa Cruz Biotechnology, Santa Cruz, CA). The resultant immune
complexes were immobilized by incubation with IgG cross-linked to agarose for
2 hat 4°C, centrifuged, washed, boiled for 5 min in sample buffer, and
again centrifuged. For studies with platelet-endothelial cell adhesion
molecule (PECAM)-1, a rabbit polyclonal antibody raised against bovine PECAM-1
(generously provided by Dr. S. M. Albelda, University of Pennsylvania Medical
School, Philadelphia, PA) and protein G cross-linked to agarose (Sigma) were
used. The supernates were then processed for immunoblotting with
antiphosphotyrosine (4G10) antibody as described above. To control for
discrepancies in immunoprecipitation and/or loading efficiencies, we stripped
and reprobed blots with the immunoprecipitating antibody. The blots were
subsequently incubated with HRP-conjugated anti-mouse IgG (Transduction
Laboratories) or HRP-conjugated anti-goat IgG (Santa Cruz) and developed with
ECL. Blots were scanned by laser densitometry, and the
phosphotyrosine-containing bands were normalized to the immunoprecipitated
protein of interest.
Coimmunoprecipitation assays. EC were lysed with ice-cold lysis
buffer containing 50 mM Tris · HCl (pH 7.4), 1% Triton X-100, 150 mM
NaCl, 1 mM phenylmethylsulfonyl fluoride, 1 mg/ml leupeptin, 1 mg/ml pepstatin
A, 1 mg/ml aprotinin, 100 mg/ml type I DNase, 1 mM vanadate, 1 mM sodium
fluoride, 10 mM disodium pyrophosphate, 500 µM paranitrophenol, and 1 mM
PAO, passed through a 25-gauge needle several times, and processed as
described above. Lysates of EC treated with vanadate or media alone were
precleared by incubation with anti-goat IgG cross-linked to agarose (Sigma)
for 1 h at 4°C and then incubated overnight at 4°C with a goat
polyclonal antibody raised against VE-cadherin (Santa Cruz). The resultant
immune complexes were processed as described above and immunoblotted with
specific murine monoclonal antibodies raised against -,
-,
p120-catenin (Transduction Laboratories) or actin (Amersham). The blots were
subsequently incubated with HRP-conjugated anti-mouse IgG (Transduction
Laboratories) and developed with ECL. To control for efficiency of
immunoprecipitation and protein loading and transfer, we stripped and reprobed
blots with the anti-VE-cadherin antibody. The blots were scanned by laser
densitometry, and the catenin of interest was normalized to VE-cadherin.
Cloning and expression of a glutathione S-transferase fusion protein
for the cytoplasmic domain of human VE-cadherin. We isolated
poly(A)+ mRNA from confluent human pulmonary artery EC using an
mRNA isolation kit (Ambion, Austin, TX), and we synthesized the cDNA using
oligo dT primers and an avian myeloblastosis virus reverse transcriptase
according to the manufacturers protocols (cDNA kit, Ambion). The cytoplasmic
domain of VE-cadherin (nt 1,8852,379; accession number X79981
[GenBank]
) was
cloned by PCR amplification from the reversed transcribed cDNA using a
5' forward primer 5'-GCGTCG G G A T C C C C C G G C G G C G G C T
C C G G A A G-3' that introduced a BamHI restriction site and a
3' reverse primer 5'-G C G G T C G A A T T C T A A T A C A G C A G
C T C C T C C C G-3' that introduced a 3' EcoRI
restriction site into the amplified VE-cadherin cDNA. The PCR products were
subcloned into pGEX-5X-1 expression vector (Amersham Pharmacia, Piscataway,
NJ), and the sequences were confirmed by DNA sequencing. Constructs for the
cytoplasmic domain of VE-cadherin and the glutathione S-transferase
(GST) (vector) alone were introduced into BL-21 Gold cells (Stratagene) and
expressed by induction with 1 mM isopropyl
-D-thiogalactopyranoside. GST-fusion proteins were purified
with the B-Per Kit (Pierce, Rockford, IL) according to manufacturer's
protocols.
GST-VE-cadherin binding assays. GST fusion protein containing the
cytoplasmic domain of human VE-cadherin was preadsorbed to glutathione
Sepharose 4B beads (Pharmacia) and then incubated with EC lysates. The
VE-cadherin-binding EC proteins bound to the beads were extensively washed,
boiled in sample buffer, resolved by SDS-PAGE, transferred to PVDF, and probed
with antibodies raised against either - or
-catenin. Simultaneous
GST/bead controls also were performed.
Human neutrophil preparation. Whole peripheral blood from healthy human volunteers was collected into acid citrate dextran (Sigma) solution, and neutrophils were isolated by dextran erythrocyte sedimentation and density gradient centrifugation through Ficoll-Hypaque (Sigma) as previously described (30). Neutrophils were resuspended at 107 cells/ml in Hanks' balanced salt solution without divalent cations (HBSS) (Life Technologies, Gaithersburg, MD) and incubated with 5 µM calcein AM (Molecular Probes) for 40 min with gentle agitation in the dark (3). Neutrophils were washed three times with HBSS- after which their purity was >95% by differential counts of Wright-stained smears and viability >98% by trypan blue dye exclusion.
Assay for transendothelial neutrophil migration. To establish the functional integrity for each monolayer, we performed transendothelial [14C]BSA flux across EC cultured to postconfluence on gelatin-impregnated polycarbonate filters (13-mm diameter, 3-µM pore size) as described in Assay of transendothelial albumin flux. Only EC monolayers retaining >97% of the [14C]BSA tracer were studied. The monolayers were then exposed to vanadate-enriched medium or to medium alone in the presence or absence of the PTK inhibitor herbimycin A (1.0 µM, Sigma), which was introduced 16 h before and throughout incubation with vanadate or medium alone. The treated monolayers were inserted into wells containing IL-8 (3 µM) or media alone. Calcein AM-labeled neutrophils (5 x 105 cells/well) were introduced into the upper compartments of assay chambers and incubated for 2 h at 37°C. Each lower compartment was sampled and fluorometrically assayed. After migration through EC monolayers cultured on filters, >99% of fluorescence remained neutrophil-associated (data not shown). A standard curve was established for each experiment from which neutrophil numbers could be interpolated from fluorescence units, and transendothelial migration of neutrophils was expressed as a percentage of total cells that migrated.
Statistical methods. Analysis of variance was used to compare the mean responses among experimental and control groups for all experiments. Dunnett's and Scheffé's F-test were used to determine between which groups significant differences existed. A P value of <0.05 was considered significant.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Effect of PTP inhibition on EC injury. A 51Cr release
assay was used to determine whether exposures to the same concentrations and
exposure times for the PTP inhibitors, vanadate and PAO, that induce
endothelial barrier dysfunction also might induce EC injury. The
51Cr release assay detects defects in the plasma membrane that
permit passage of molecules 1,000 Da. EC monolayers preloaded with
51Cr were exposed for 6h to vanadate (25 µM), PAO (0.5 µM),
or media alone. Mean (± SE) 51Cr release from either
vanadateor PAO-exposed EC (13.07 ± 0.35%, n = 10, and 12.89
± 0.47%, n = 10, respectively) was not significantly different
from release from the simultaneous media controls (12.32 ± 0.51%,
n = 10). These studies indicate that changes in barrier function in
response to PTP inhibition cannot be ascribed to EC injury.
PTP inhibition promotes intercellular gap formation in postconfluent EC monolayers and preformed EC tubes. We questioned whether the vanadate- and PAO-induced loss of barrier function could be explained through opening of the paracellular pathway. Accordingly, EC monolayers exposed for 6 h to media alone (Fig. 2A), 0.5 µM PAO (Fig. 2B), or 25 µM vanadate (Fig. 2C) were stained with fluorescein-phalloidin, an F-actin-specific reagent. By fluorescence microscopy, EC monolayers incubated with media alone exhibited continuous transcytoplasmic actin filaments and cell-to-cell apposition without intercellular gaps (Fig. 2A). Exposure of EC to 0.5 µM PAO (Fig. 2B) or 25 µM vanadate (Fig. 2C) induced disruptions within the F-actin lattice exclusively at the EC-EC interface and circumferential redistribution of F-actin staining to the cell periphery. In other experiments, EC were cultured in Matrigel-coated wells. EC attached within 0.5 h and formed interconnected tubular networks between 6 and 16 h. PTP inhibition with either PAO (Fig. 2E) or vanadate (Fig. 2F) disrupted EC-EC interactions, whereas EC-matrix attachments were sustained (Fig. 2, DF). These data indicate that PTP inhibition with either vanadate or PAO promotes actin reorganization and intercellular gap formation in postconfluent EC monolayers as well as in preformed EC tubular structures.
|
PTP inhibition increases tyrosine phosphorylation of EC proteins. PTP inhibition with pervanadate (vanadate complexed with H2O2) has previously been demonstrated to induce rapid and dramatic increases in the tyrosine phosphorylation of proteins localized to the intercellular boundaries (4, 53). We therefore asked whether vanadate-induced intercellular gap formation and increases in transendothelial [14C]BSA flux could be correlated to increases in protein tyrosine phosphorylation. Accordingly, EC exposed for 1 h to increasing concentrations of vanadate (0.525 µM) were processed for phosphotyrosine immunoblotting (Fig. 3). Vanadate at concentrations as low as 0.5 µM increased the phosphotyrosine signal compared with the effect seen with media alone. This same vanadate concentration was the lowest concentration that induced increments in [14C]BSA flux (Fig. 1A). PTP inhibition with vanadate (25 µM) induced tyrosine phosphorylation of a number of EC proteins including several that migrated with an apparent Mr of 240,000, 220,000, 185,000, 165,000, 135,000, 110,000, 95,000, and 66,000. Over the range of 2.525 µM vanadate, this increase in phosphotyrosine signal was concentration dependent. At a fixed concentration of vanadate (25 µM), exposure times as brief as 5 min were associated with increases in phosphotyrosine signal compared with the simultaneous media control (Fig. 3B). These vanadate-induced increases in tyrosine phosphorylation precede the first demonstrable increases in transendothelial [14C]BSA flux (see Fig. 1C).
|
Immunolocalization of phosphotyrosine-containing proteins to the EC-EC boundaries with PTP inhibition. To determine the subcellular localization of phosphotyrosine-containing proteins in vanadate-treated EC, postconfluent monolayers exposed to vanadate or media alone were probed with a FITC-conjugated antiphosphotyrosine antibody and analyzed by epifluorescence microscopy (Fig. 4). EC incubated with vanadate (25 µM) displayed a fluorescence signal that was predominately restricted to the intercellular boundaries (Fig. 4, BD), whereas the cells provided with media alone lacked this pattern of immunofluorescence (Fig. 4A). This phosphotyrosine signal was evident as early as 10 min (Fig. 4B) with dramatic increases observed by 1 h (Fig. 4D). These studies indicate that PTP inhibition with vanadate promotes tyrosine phosphorylation of proteins that are either enriched to, or upon, phosphorylation translocate to the EC-EC junctions. Furthermore, these tyrosine phosphorylation events are temporally proximal to intercellular gap formation (Fig. 2, C and E) and opening of the paracellular pathway (Fig. 1C).
|
PTP inhibition increases tyrosine phosphorylation of EC-EC junctional
proteins. To assess whether PTP inhibition with vanadate increases
tyrosine phosphorylation of EC-EC adherens junction (ZA) proteins, EC exposed
for 1 h to vanadate (25 µM) or media alone were immunoprecipitated with
anti-VE-cadherin, anti--catenin, anti-
-catenin, or
anti-p120ctn antibodies, and the immunoprecipitates were processed
for phosphotyrosine immunoblotting (Fig.
5A). PTP inhibition with vanadate increased tyrosine
phosphorylation of VE-cadherin
5-fold,
-catenin
4-fold,
-catenin
4-fold, and p120ctn
5-fold compared with
media controls. These data indicate that vanadate, at a concentration and
exposure time that opens the paracellular pathway (see Figs.
1, A and C,
and 2C), increases
tyrosine phosphorylation of components of the ZA multiprotein complex. In
similar experiments, vanadate also increased tyrosine phosphorylation of
PECAM-1
4-fold (Fig.
5B). Whether tyrosine phosphorylation of one or more of
these junctional proteins regulates EC-EC adhesion in the context of
paracellular pathway function and/or angiogenesis is unclear.
|
Effect of PTP inhibition on the state of ZA assembly. Increased
tyrosine phosphorylation of ZA proteins can disrupt the intercellular
cadherin-catenin linkage and/or uncouple the ZA from the actin cytoskeleton
(6,
34,
39). Thus we examined whether
vanadate-induced tyrosine phosphorylation of ZA protein components altered the
association of VE-cadherin with -,
-, or p120-catenin, or with
the actin cytoskeleton. Immunoprecipitation of VE-cadherin from lysates of EC
treated for 1 h with vanadate or media alone coimmunoprecipitated both
catenins and actin, and no differences in coimmunoprecipitation of either the
catenins or actin could be demonstrated
(Fig. 6A).
Immunoprecipitation with an irrelevant antibody did not precipitate
VE-cadherin,
-,
-, and p120-catenin, or actin (data not shown).
To confirm our findings that the state of tyrosine phosphorylation of either
- or
-catenin does not influence its ability to bind VE-cadherin,
lysates from EC exposed for 1 h to vanadate (25 µM) or media alone were
incubated with a GST fusion protein containing the cytoplasmic domain of human
VE-cadherin coupled to glutathione Sepharose beads or a GST bead control
(Fig. 6B). The
VE-cadherin-binding proteins bound to the beads were extensively washed and
processed for immunoblotting with either anti-
-catenin or
anti-
-catenin antibodies. In the presence or absence of PTP inhibition,
the binding of either
- or
-catenin to the cytoplasmic domain of
VE-cadherin was equivalent (Fig.
6B). These combined data
(Fig. 6, A and
B) suggest that in EC, tyrosine phosphorylation of ZA
protein components does not dictate disassembly of the ZA multiprotein complex
or uncoupling of the ZA from the actin cytoskeleton but is sufficient to
promote intercellular gap formation and opening of the paracellular
pathway.
|
Effect of PTP inhibition on colocalization of VE-cadherin with
either - or
-catenin. To extend our in
vitro findings to an intact cell system, we applied double-label
colocalization fluorescence microscopy. EC monolayers were incubated for
16 h with vanadate (25100 µM) or media alone, after which
they were probed with antibodies against VE-cadherin and either
- or
-catenins and processed for epifluorescence microscopy
(Fig. 7). In the media control
monolayers, discrete and continuously linear staining for VE-cadherin
(Fig. 7, A and
C) and either
-catenin
(Fig. 7A) or
-catenin (Fig.
7C) was evident. These fluorescent signals were almost
exclusively restricted to intercellular boundaries and revealed a high degree
of VE-cadherin-catenin colocalization (Fig.
7, A and C). In some areas, these same control
cells also exhibited short, discontinuous regions of EC-EC contact also
displaying a high degree of cadherin-catenin colocalization
(Fig. 7, A and
C). PTP inhibition with increasing concentrations of
vanadate for increasing exposure times induced both dose- and time-dependent
changes, the most subtle of which were seen after a 1-h exposure to 25 µM
vanadate, where no changes in colocalization of VE-cadherin with either
-/
-catenin could be detected (data not shown). The most dramatic
changes were evident after a 6-h exposure to 100 µM vanadate
(Fig. 7, B and
D). In these EC, PTP inhibition was associated with a
marked increase in the arrays of multiple short segments of cadherin/catenin
staining that retained a remarkable degree of VE-cadherin/catenin
colocalization (Fig. 7, B and
D). Vanadate also induced intercellular gaps around which
both cadherin and catenin staining was markedly diminished
(Fig. 7, B and
D). This decrease in VE-cadherin and
-/
-catenin staining at EC-EC boundaries was not associated with
an increase in cytoplasmic staining of these same ZA proteins
(Fig. 7, B and
D). Even in the areas where VE-cadherin and catenin
staining was less abundant, their colocalization was sustained. Together,
these data indicate that although PTP inhibition and its attendant tyrosine
phosphorylation of ZA proteins promote profound reorganization of EC-EC
contacts, VE-cadherin and
- and
-catenins continue to
colocalize.
|
PTK inhibition protects against vanadate-induced protein tyrosine
phosphorylation and opening of an endothelial paracellular pathway. We
previously have demonstrated that PTK inhibition protects against
agonist-induced protein tyrosine phosphorylation, intercellular gap formation,
and opening of an endothelial paracellular pathway
(7,
31,
61). To determine whether
PTK-dependent phosphorylation of EC proteins was operative during barrier
dysfunction in response to PTP inhibition, we employed the PTK inhibitor
herbimycin A (Fig.
8A). The mean (± SE) pretreatment baseline barrier
function was 0.010 ± 0.001 pmol/h (n = 72), and there were no
significant differences among the experimental groups. [14C]BSA
flux across EC monolayers treated with herbimycin A (1.0 µM) alone was not
different from that of the media controls. A 6-h vanadate (25 µM) exposure
increased transendothelial [14C]BSA flux and pretreatment of EC
monolayers with herbimycin A protected against this increment by 80%. To
confirm that this concentration of herbimycin A that protected against loss of
barrier function also diminished tyrosine phosphorylation of EC proteins, we
processed EC exposed to vanadate with or without herbimycin A for
phosphotyrosine immunoblotting (Fig.
8B). PTK inhibition with herbimycin A diminished
vanadate-induced tyrosine phosphorylation of EC proteins. Thus the PTK
inhibitor that protected against opening of the paracellular pathway gained
entry into EC and blocked vanadate-induced protein tyrosine phosphorylation.
The ability of prior PTK inhibition with herbimycin A to block
vanadate-induced increments in albumin flux demonstrates that ongoing,
constitutive protein tyrosine phosphorylation is required for opening of the
paracellular pathway in response to PTP inhibition. These data indicate that
both PTKs and PTPs actively regulate the endothelial paracellular pathway.
|
PTP inhibition promotes transendothelial migration of neutrophils.
To determine whether this same tyrosine phosphorylation-responsive
paracellular pathway might be accessed by migrating neutrophils, we employed
an assay for transendothelial migration of neutrophils. PTP inhibition with
vanadate enhanced transendothelial migration of neutrophils 3.5-fold
compared with the simultaneous media controls
(Fig. 8C). Prior PTK
inhibition with herbimycin A protected against the vanadate effect by >80%.
These data suggest that transendothelial paracellular migration of neutrophils
is also regulated through the action of both EC PTKs and PTPs.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Tyrosine phosphorylation of ZA proteins reduces EC-EC junctional integrity
and opens the endothelial paracellular pathway
(7,
20,
31,
61). Although most studies on
the regulation of this tyrosine phosphorylation-responsive paracellular
pathway have emphasized PTK-driven events
(7,
8,
20,
33,
34,
37,
54), evidence also exists for
its counterregulation by PTPs
(53,
57). As cells achieve
confluence with the formation and stabilization of intercellular junctions,
PTP activity increases (25,
50), whereas ZA protein
tyrosine phosphorylation decreases
(4,
43). The Meprin/A5/PTPµ
domain-containing receptor PTPs whose ectodomains homophilically interact,
including PTP-µ, PTP-, and PTP-
, become trapped at the
cell-cell interface (10,
15,
23,
28). Here, their catalytic
domains are restrained in close proximity to EC-EC junctions, including the
ZA. The cytoplasmic domains of these same three PTPs as well as another
receptor PTP, LAR-PTP, and two nonreceptor PTPs, SHP-2 and PTP-1B, each bind
to and/or dephosphorylate ZA proteins
(6,
11,
12,
15,
23,
42,
47,
55,
62). VE-PTP reportedly
regulates the phosphorylation state of VE-cadherin independently of its
catalytic PTP domain (48).
Established mediators of vascular permeability and angiogenesis, including
TNF-
, IL-1, and thrombin, each perturb PTP catalytic activity
(2,
29,
32,
55). Finally, PTP inhibition
itself increases tyrosine phosphorylation of proteins almost exclusively
restricted at cell-cell boundaries
(53) and decreases both
cell-cell junctional integrity
(57) and transcellular
electrical resistance (53).
With these combined data in mind, we asked whether PTPs might participate in
the regulation of endothelial barrier function.
In our studies, nonselective PTP inhibition with either of two structurally and functionally dissimilar agents, vanadate and PAO, increased transendothelial albumin flux in a dose- and time-dependent manner (Fig. 1). F-actin fluorescence microscopy of postconfluent EC monolayers subjected to PTP inhibition demonstrated intercellular gap formation with circumferential redistribution of F-actin (Fig. 2, AC). Vanadate-sensitive PTPs that target cytoskeletal proteins (59) could participate in the actin reorganization seen in EC after PTP inhibition. Similarly, PTP inhibition induced intercellular gap formation in preformed EC tubes (Fig. 2, DF). These data indicate that nonselective PTP inhibition decreases endothelial barrier function at the level of the paracellular pathway.
To increase our understanding of PTP(s) that may regulate the paracellular
pathway, we pursued their substrates. After PTP inhibition, phosphotyrosine
immunoblotting demonstrated phosphotyrosine-containing bands, several of which
migrated with gel mobilities compatible with one or more EC-EC junctional
proteins. The dose and time requirements for increased tyrosine
phosphorylation in response to PTP inhibition were compatible with those
necessary for loss of barrier function. Phosphotyrosine fluorescence
microscopy immunolocalized these phosphotyrosine-containing proteins almost
exclusively to EC-EC boundaries. This suggested that PTPs dephosphorylate
proteins associated with intercellular junctions and/or proteins that upon
tyrosine phosphorylation translocate to the cell periphery. The receptor PTPs
that sequester at the cell-cell interface through ectodomain homophilic
adhesion (10,
15,
23,
28) might contribute to this
phosphotyrosine signal preferentially displayed within intercellular
boundaries in response to PTP inhibition
(Fig. 4). When we
immunoscreened for ZA protein substrates, the tyrosine phosphorylation states
of VE-cadherin and -,
-, and p120 catenins each were increased.
It is conceivable that one or more of the reported ZA-associated PTPs,
especially two expressed in EC, PTP-µ and SHP-2
(9,
55), are involved. In another
study, PTP inhibition in bovine brain EC increased tyrosine phosphorylation of
the tight junctional proteins zonula occludins-1 and -2
(53). It is possible that
ZA-associated PTPs bind to and/or dephosphorylate substrates within other
junctional complexes that reside within the same subcellular compartment. For
example, SHP-2, which is recruited to tyrosine phosphorylated PECAM-1, also
restrains tyrosine phosphorylation of ZA proteins
(55).
The mechanism(s) by which the tyrosine phosphorylation state of one of more
ZA proteins can, through in-to-out signaling, regulate VE-cadherin
ectodomain-mediated homophilic adhesion is poorly understood. Because ZA
disassembly and disruption of the ZA-actin cytoskeletal linkage is known to
reduce cell-cell homophilic adhesion
(34,
39), we asked whether PTP
inhibition might open the endothelial paracellular pathway through uncoupling
of VE-cadherin from its catenin binding partners and the actin cytoskeleton.
PTP inhibition with vanadate (25 µM) for 1 h, which is associated with
dramatic increases in both ZA protein tyrosine phosphorylation
(Fig. 5A) and
paracellular permeability (Fig.
1C), did not decrease VE-cadherin colocalization with
either - or
-catenin (data not shown). Similarly, this same PTP
inhibition did not, under moderately stringent conditions, diminish
coimmunoprecipitation of
-,
-, or p120 catenin or actin with
VE-cadherin (Fig. 6A).
In an in vitro binding assay, no decrease in binding of either
tyrosine-phosphorylated
- or
-catenin to immobilized
nonphosphorylated VE-cadherin could be detected
(Fig. 6B), suggesting
that the tyrosine phosphorylation state of VE-cadherin is not critical to
VE-cadherin-catenin binding. PTP inhibition with vanadate only at higher
concentrations for prolonged exposure times was associated with diminished
VE-cadherin/catenin staining at intercellular boundaries
(Fig. 7). In these monolayers,
VE-cadherin and
-/
-catenins could be localized to short,
discrete, linear segments (Fig. 7,
B and D). Even after this more rigorous PTP
inhibition (i.e., vanadate 100 µM, 6 h), VE-cadherin colocalization with
either
-/
-catenin was remarkably sustained. It is conceivable
that these discrete, discontinuous segments of VE-cadherin-catenin
colocalization represent an intermediate state for ZA organization that allows
for increased flexibility as EC populations transition from a quiescent,
contact-inhibited barrier to migratory EC responding to intimal injury or EC
clusters organizing into tubes during angiogenesis. These combined data
demonstrate that tyrosine hyperphosphorylation of VE-cadherin or any of its
associated catenins does not in itself dictate VE-cadherin-catenin
disassembly.
In postconfluent EC, membrane-spanning VE-cadherin molecules are trapped at
EC-EC junctions through ectodomain homophilic adhesion. Opening of the
paracellular pathway theoretically permits disengagement of homophilically
bound VE-cadherin ectodomains and lateral mobility in the lipid bilayer with
redistribution across the plasma membrane. It is conceivable that as
VE-cadherin and either -or
-catenin leave the intercellular
junctions, they do so together as a VE-cadherin-catenin complex. This would
permit more dynamic and efficient ZA disassembly/reassembly in response to
rapidly changing physiological demands. The EC response to PTP inhibition was
not uniform across the entire EC monolayer. Our VE-cadherin
coimmunoprecipitation and GST-VE-cadherin in vitro binding assays reflect only
the average of the cellular events occurring throughout the entire monolayer
and might not detect subtle changes in protein-protein interactions restricted
to an EC subpopulation and/or to a subcellular compartment. Other studies with
non-EC systems have generated conflicting results. In several studies,
tyrosine phosphorylation of ZA proteins, especially
-catenin, is
associated with disruption of the cadherin-actin cytoskeletal linkage and ZA
disassembly (34,
39). In other studies,
including our own findings, tyrosine phosphorylation of ZA proteins did not
appear to promote ZA disassembly or disruption of the cadherin actin
cytoskeleton linkage (8,
20). It is well known that
-catenin couples VE-cadherin not only to the actin cytoskeleton, but to
the intermediate filament network as well
(41,
56). In a recent study,
histamine, an established mediator of microvascular paracellular permeability,
decreased VE-cadherin coimmunoprecipitation of the intermediate filament
protein vimentin (52).
Therefore, under certain conditions of increased paracellular permeability,
VE-cadherin disengages from the intermediate filament network, whereas
VE-cadherin-actin association is sustained. The current study indicates that
dramatic changes in the state of ZA assembly and its linkage to the actin
cytoskeleton are not prerequisites for tyrosine phosphorylation-dependent
opening of the endothelial paracellular pathway.
In our studies, EC were subjected to PTP inhibition in the presence of serum and whatever mitogenic growth factors it contains. We therefore asked whether a basal level of ongoing protein tyrosine phosphorylation was required for the paracellular pathway to open in response to PTP inhibition. Prior PTK inhibition with a broad-spectrum PTK inhibitor blocked the PTP effect. Therefore, constitutively tyrosine-phosphorylated proteins may be a prerequisite for EC responsiveness to PTP inhibition. Our studies confirm the importance of both PTKs and PTPs in endothelial paracellular pathway regulation. PTPs appear to restrain PTK-driven protein tyrosine phosphorylation, whereas PTK-driven tyrosine phosphorylation appears to prime and maintain EC in a tonic state of lowered threshold for PTP inhibition.
The same tyrosine phosphorylation-responsive endothelial paracellular
pathway also could be accessed by neutrophils. An EC-EC junctional protein
that participates in transendothelial polymorphonuclear neutrophils (PMN)
migration, PECAM-1 (or CD31), is a substrate for tyrosine phosphorylation
(21,
45,
49). Vanadate increased both
tyrosine phosphorylation of PECAM-1 (Fig.
5B) and PMN migration
(Fig. 8C), whereas
herbimycin A blocked both PECAM-1 phosphorylation (data not shown) and PMN
migration. Whether there is a causal relationship between the tyrosine
phosphorylation state of PECAM-1 and transendothelial PMN migration is not
known. The tyrosine phosphorylation state of PECAM-1 changes with EC adhesion
and migration (45) and
regulates its homophilic adhesion
(21), its ability to serve as
a reservoir for tyrosine-phosphorylated -catenin
(35), and its recruitment of
SH-2 containing PTPs including SHP-2
(36). Although PECAM-1 employs
distinct EC-EC adhesion mechanisms from the ZA
(5), it may serve as a
scaffolding structure to recruit and position SHP-2 in close proximity to ZA
protein substrates.
Tyrosine phosphorylation of ZA protein components and opening of the endothelial paracellular pathway can be induced by many agonists through various cell surface receptors. To circumvent these multiple signaling events and to study the role of PTPs in regulation of the endothelial paracellular pathway, we used pharmacological agents that inhibit PTP activity. In this study, we demonstrate that PTP inhibition alone promotes tyrosine phosphorylation of ZA proteins coincident with increases in the movement of macromolecules and neutrophils across the endothelium barrier. Furthermore, our studies indicate that dramatic changes in the state of ZA assembly and its linkage to the actin cytoskeleton are not prerequisites for opening of the paracellular pathway. It is conceivable and even likely that phosphoproteins other than ZA components are operative during the EC response to PTP inhibition. These findings suggest that PTP-dependent modification of ZA protein components may be a vital mechanism employed to regulate movement of macromolecules and neutrophils across the endothelium as well as stability of EC capillary tubes.
![]() |
ACKNOWLEDGMENTS |
---|
This work was supported in part by the Office of Research and Development, Department of Veterans Affairs; National Heart, Lung, and Blood Institute Grants HL-63217 and HL-70155; and the American Heart Association.
![]() |
FOOTNOTES |
---|
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.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Visit Other APS Journals Online |