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(American Journal of Pathology. 2005;166:1247-1258.)
© 2005 American Society for Investigative Pathology

Receptor Protein Tyrosine Phosphatase µ Regulates the Paracellular Pathway in Human Lung Microvascular Endothelia

Xiu Fen Sui*, Timothy D. Kiser*, Sang Won Hyun*, Daniel J. Angelini*, Robert L. Del Vecchio{dagger}, Bradford A. Young*, Jeffrey D. Hasday*, Lewis H. Romer{ddagger}, Antonino Passaniti*, Nicholas K. Tonks{dagger} and Simeon E. Goldblum*

From the Departments of Medicine and Pathology,* Division of Infectious Diseases and Pulmonary Medicine, Mucosal Biology Research Center, University of Maryland School of Medicine, Baltimore, Maryland; Cold Spring Harbor Laboratory,{dagger} Cold Spring, New York; and the Departments of Medicine, Anesthesiology, Cell Biology, and Pediatrics,{ddagger} The Johns Hopkins University School of Medicine, Baltimore, Maryland


   Abstract
Top
 Abstract
Materials and Methods
 Results
 Discussion
 References
 
The pulmonary vascular endothelial paracellular pathway and zonula adherens (ZA) integrity are regulated, in part, through protein tyrosine phosphorylation. ZA-associated protein tyrosine phosphatase (PTP)s are thought to counterregulate tyrosine phosphorylation events within the ZA multiprotein complex. One such receptor PTP, PTPµ, is highly expressed in lung tissue and is almost exclusively restricted to the endothelium. We therefore studied whether PTPµ, in pulmonary vascular endothelia, associates with and/or regulates both the tyrosine phosphorylation state of vascular endothelial (VE)-cadherin and the paracellular pathway. PTPµ was expressed in postconfluent human pulmonary artery and lung microvascular endothelial cells (ECs) where it was almost exclusively restricted to EC-EC boundaries. In human lung microvascular ECs, knockdown of PTPµ through RNA interference dramatically impaired barrier function. In immortalized human microvascular ECs, overexpression of wild-type PTPµ enhanced barrier function. PTPµ-VE-cadherin interactions were demonstrated through reciprocal co-immunoprecipitation assays and co-localization with double-label fluorescence microscopy. When glutathione S-transferase-PTPµ was incubated with purified recombinant VE-cadherin, and when glutathione S-transferase-VE-cadherin was incubated with purified recombinant PTPµ, PTPµ directly bound to VE-cadherin. Overexpression of wild-type PTPµ decreased tyrosine phosphorylation of VE-cadherin. Therefore, PTPµ is expressed in human pulmonary vascular endothelia where it directly binds to VE-cadherin and regulates both the tyrosine phosphorylation state of VE-cadherin and barrier integrity.

The pulmonary vascular endothelium presents a selective barrier that actively regulates paracellular movement of circulating macromolecules and cells into extravascular tissues and compartments.1,2 We3,4 and others5-8 have demonstrated that the endothelial paracellular pathway through which macromolecules flux is regulated, in part, through protein tyrosine phosphorylation. Established mediators of vascular permeability, including endotoxin and counteradhesive proteins, increase tyrosine phosphorylation of endothelial cell (EC) proteins almost exclusively restricted to the intercellular boundaries whereas prior protein tyrosine kinase (PTK) inhibition protects against opening of the paracellular pathway in response to these same agonists.3,4 Several substrates for agonist-induced tyrosine phosphorylation have been identified as components of the EC-EC adherens junction or zonula adherens (ZA),4,6 an intercellular junctional complex that modulates homophilic cell-cell adhesion.1,9-11

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 in ECs.9,10 Although multiple cadherins can be co-expressed and differentially distributed in ECs,12 VE-cadherin appears to be unique to ECs and is localized to their intercellular junctions.13 Further, anti-VE-cadherin antibodies increase transendothelial flux of macromolecules and established mediators of EC injury alter VE-cadherin distribution. At least three cytoplasmic proteins, collectively termed catenins, form multiprotein complexes that participate in anchoring the cytoplasmic domain of cadherins to actin microfilaments.14-16 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.1,11

The state of ZA protein tyrosine phosphorylation is central to the regulation of the ZA-actin cytoskeletal linkage and homophilic cell-cell adhesion17-19 and, as we3,4 and others5,6 have shown, to the maintenance of endothelial barrier function. More specifically, the tyrosine phosphorylation state of VE-cadherin appears to influence the endothelial paracellular pathway.6,20 Whether tyrosine phosphorylation of its cytoplasmic domain directly regulates homophilic interactions between its ectodomains is unclear. Protein tyrosine phosphatases (PTPs) are thought to play a crucial role in regulating the state of ZA protein tyrosine phosphorylation and assembly and endothelial barrier function. Increased expression of a number of PTPs parallels increases in cell density.21-24 In contact-inhibited confluent human umbilical vein ECs (HUVECs), membrane-associated PTP activity is increased ~10-fold compared to subconfluent ECs.22,24 In postconfluent pulmonary vascular ECs, we have found that for some mediators, rigorous PTP inhibition is required for a consistent, reproducible, agonist-induced phosphotyrosine signal.4 Esser and colleagues6 reported similar findings in vascular endothelial growth factor-stimulated HUVECs. Further, co-administration of PTP inhibitors with some of these same agonists, at concentrations that alone do not alter barrier function, enhances mediator-induced increments in transendothelial albumin flux.3,4 Several such agonists have been shown to alter PTP expression or activity25-27 and/or PTP-substrate interactions.7 Finally, PTP inhibition itself induces dose- and time-dependent increments in protein tyrosine phosphorylation, intercellular gap formation, and loss of barrier function both in vitro5,28 and in vivo.8 The phosphotyrosine-containing proteins are immunolocalized predominantly to the intercellular boundaries and several have been identified as ZA proteins.5,28 These combined data suggest that pulmonary vascular ECs express PTPs that associate with and dephosphorylate ZA and possibly other intercellular junctional proteins. In fact, a growing number of ZA-associated PTPs have been demonstrated in various epithelia and other tissues.29-36 The receptor PTPs, PTPµ,23,37,38 PTPK,32 PTP{lambda},33 VE-PTP, also known as PTPß,35 density-enhanced phosphatase (DEP)-1,36,39 and a member of the leukocyte common antigen-related protein (LAR)-PTP family,34 each have been shown to bind to and/or dephosphorylate ZA proteins. In HUVECs, the SH2 domain-containing nonreceptor PTP, SHP-2, binds to ß-catenin and restrains phosphorylation of ß-, {gamma}-, and p120 catenins.7 In chick retinal tissue, another nonreceptor PTP, PTP1B, associates with N-cadherin and dephosphorylates ß-catenin.19 It appears that one or more PTPs, possibly in concert, regulate tyrosine phosphorylation events within the ZA multiprotein complex of various cells.

The in vivo tissue expression of these ZA-associated PTPs are distinct; some are ubiquitous whereas others are restricted.40,41 One such PTP, PTPµ, is highly expressed in lung,37,41 is almost exclusively restricted to the vascular endothelium,41,42 and is co-localized with flk-1, a murine EC-specific receptor for vascular endothelial growth factor, with high expression in the pulmonary vasculature. In other in vivo studies of adult rat and porcine tissues, PTPµ can be immunolocalized almost exclusively to vascular EC-EC junctions where it co-localizes with cadherin-catenin complexes.42 These combined studies suggest that PTPµ may be relevant to the regulation of the tyrosine phosphorylation-responsive pulmonary vascular endothelial paracellular pathway.

PTPµ is a multidomain and apparently multifunctional protein that may influence the endothelial paracellular pathway through one or more mechanisms. The extracellular domain of this 195-kd protein contains in tandem an N-terminal MAM (Meprins A and B, Xenopus A5 glycoprotein, PTPmu) domain, an immunoglobulin (Ig)-like repeat, and four fibronectin III-like repeats.40 The ectodomain requires its Ig-like repeat to participate in homophilic adhesion with an identical molecule on neighboring cells.40,43 The intracellular segment contains two catalytic domains, the first or NH2-terminal of which is active.40 A single (C->S) amino acid mutation within the first catalytic domain renders PTPs catalytically inactive and such mutants can act as dominant-negative molecules.44 Double (Y->F, D->A) amino acid mutations within the same catalytic domain result in a substrate-trapping, catalytically impaired mutation that might retain low residual PTP activity.45,46 Such mutations impair catalysis without affecting affinity for substrates, thereby stabilizing PTPµ-substrate interactions and potentially interfering with downstream signaling events. In PTPµ, the ~70-amino acid juxtamembranous segment between the membrane-spanning domain and first catalytic site shares ~20% amino acid identity with the conserved, cytoplasmic portion of cadherin.37,38,40 This segment interacts with both phosphatase domains and may regulate catalytic activity.47 In the extracts of rat lung and various cell lines, PTPµ interacts with the classical cadherins, E-cadherin, N-cadherin, and cadherin-430 and possibly one or more of the catenins.31 Among the classical cadherins, the VE-cadherin cytoplasmic domain is the least conserved.48 Whether PTPµ can similarly bind to the vascular EC-specific cadherin, VE-cadherin, is unknown. In the current studies, we determined whether PTPµ catalytic activity regulates the human microvascular endothelial paracellular pathway. Further, we tested whether PTPµ expressed in pulmonary vascular endothelia directly/indirectly associates with and/or regulates the tyrosine phosphorylation state of VE-cadherin.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
EC Culture

Human pulmonary artery and lung microvascular ECs (Clonetics Corp., San Diego, CA) were cultured in EC growth medium (EBM-2, Clonetics) containing 5% fetal bovine serum (Hyclone Laboratories, Logan, UT), human recombinant epidermal growth factor, human recombinant insulin-like growth-factor-1, human basic fibroblast growth factor, vascular endothelial growth factor, hydrocortisone, ascorbic acid, gentamicin, and amphotericin B. Only ECs at passages 2 to 7 were studied. HMEC-1 cells, a simian virus (SV) 40 T antigen transformed human dermal microvascular EC line (CDC, Atlanta, GA), were also cultured with EBM-2 in the presence of 10% fetal bovine serum.

Reverse Transcriptase (RT)-Polymerase Chain Reaction (PCR) for PTPµ

Poly(A)+ RNA was isolated from confluent human pulmonary artery ECs, human lung microvascular ECs, and HMEC-1 cells. Complementary DNA was generated from RNA using oligo(dT) primers and AMV reverse transcriptase. This cDNA was used as a template for amplification with DyNAzyme EXT DNA polymerase and primers A (5'-CGGTGCVATGGACATCCTGCC-3') and B (5'-CTTGTACTGATCCAGGAGGTC-3') that corresponded to the cytoplasmic domain of PTPµ (3652 bp to 4297 bp). The PCR was performed in a Perkin-Elmer GeneAmp PCR system 2400 (Applied Biosystems, Foster City, CA). After an initial 2 minutes of denaturation at 94°C, 30 cycles compromising 30 seconds at 94°C (denaturation), 1 minute at 60°C (annealing), 1 minute at 72°C (extension) was completed followed by a 7-minute final extension at 72°C. The purified PCR products were subcloned into a TA vector and the subcloned constructs were then expressed in XL-1 subcloning cells, isolated, purified, and sequenced to confirm the presence of PTPµ.

Immunoblotting for PTPµ and ZA Proteins

Postconfluent ECs were lysed with ice-cold lysis buffer as previously described in detail.3,4 The lysates were centrifuged, assayed for protein concentration, resolved by electrophoresis on a 6% 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 murine monoclonal antibody raised against the intracellular juxtramembranous segment of PTPµ (SK 7)49 followed by horseradish peroxidase (HRP)-conjugated rabbit anti-mouse IgG (Transduction Laboratories, Lexington, KY), and developed with enhanced chemiluminescence (ECL) (Amersham, Arlington Heights, IL). To confirm equivalent protein loading, blots were stripped with 100 mmol/L 2-mercaptoethanol, 2% SDS, 62.5 mmol/L Tris-HCl, pH 6.7, and reprobed with 0.5 mg/ml of murine anti-physarum ß-tubulin IgG2b (Boehringer-Mannheim, Indianapolis, IN) followed by HRP-conjugated anti-mouse IgG (Transduction Laboratories)3,4 and developed with ECL. In selected experiments with HMEC-1 cells, the EC lysates were processed for immunoblotting with murine monoclonal antibodies raised against the ZA proteins, VE-cadherin (cadherin-5, Transduction Laboratories), and {alpha}-, ß-, {gamma}-, and p120-catenins (Transduction Laboratories). The cadherin-5 anti-VE-cadherin antibody was raised against a peptide corresponding to a portion of the ectodomain (amino acids 26 to 194).

Immunolocalization of PTPµ and Its Co-Localization with VE-Cadherin by Epifluorescence Microscopy

For localization of PTPµ, ECs were cultured to postconfluence, washed [phosphate-buffered saline (PBS) without Ca2+/Mg2+] three times, fixed (3% paraformaldehyde at 37°C for 5 minutes), permeabilized (0.5% Triton X-100 for 5 minutes), blocked [2% bovine serum albumin (BSA)/5% horse serum (Life Technologies, Inc., Grand Island, NY) for 0.5 hours], and incubated for 1 hour with murine anti-PTPµ antibodies followed by fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse IgG (Molecular Probes Inc., Eugene, OR) as described3 with minor modifications. For co-localization of PTPµ with VE-cadherin, monolayers were probed with monoclonal SK7 anti-PTPµ antibody and goat anti-human VE-cadherin IgG (Santa Cruz Biotechnology, Santa Cruz, CA) at 1:50 dilution in 0.1% BSA in PBS followed by Cy-3-labeled goat anti-mouse IgG and FITC-conjugated donkey anti-goat IgG (3 µg/ml) (Molecular Probes Inc.). After multiple washes, the monolayers were processed for standard epifluorescence microscopy.

PTPµ Knockdown by RNA Interference

Small interfering RNA (siRNA) duplex products were designed and prepared (Dharmacon, Lafayette, CO) to target PTPµ. These duplex siRNAs included four distinct sequences GGGCAGAACUGGCCAUUAG (425 to 443) GGAAGAACGUCCUCGAAGA (1878 to 1896), CGACGAGGCUUUCUCAUUC (2406 to 2424), and GCAAUUAUAUCGAUGGUUA (2867 to 2885). An irrelevant duplex siRNA that does not correspond to any known sequence in the human genome was similarly prepared as a control. The four PTPµ siRNAs mixed together in equivalent concentrations and the control both were preincubated with TransMessenger transfection reagent (Qiagen, Valencia, CA) according to the manufacturer’s protocol and the transfection complexes presented to human lung microvascular ECs cultured to ~80% confluence for 3 hours in the absence of serum. At increasing times after transfection, ECs were lysed and processed for immunoblotting with SK-15 anti-PTPµ antibody (Oncogene, Boston, MA). To confirm equivalent protein loading, blots were stripped and reprobed with anti-ß-tubulin antibodies and developed with ECL as described above. Autoradiographs were scanned by laser densitometry (Molecular Dynamics, Sunnyvale, CA) and the PTPµ-immunoreactive bands analyzed.

Overexpression of Wild-Type (WT) PTPµ and Catalytically Impaired PTPµ in HMEC-1 Cells

A retrovirus-mediated gene transfer system (pRevTRE2; Clontech, Palo Alto, CA) was applied to stably infect HMEC-1 cells. In selected experiments, the system was tetracycline-responsive. pRP265/PTPµ38 was used as a template for overlapping PCR to generate the mutation of Asp1063 (G!T) to Ala (GCT). The resulting plasmid, pRP265/µDA, was used as the template for inverse PCR to generate the mutation of Tyr929(T!C) to Phe(TTC) to give pRP265/µY9292F,D1063A. A NgoM IV/XbaI fragment (nucleotides 2596 to 4356, PTPµ from pRP265/µYFDA was ligated to NgoM IV/XbaI digested pMT2/µ37 to produce pMT2/µYFDA. WT and YFDA PTPµ mutant constructs each were subcloned into the NotI site of pRevTRE2 (Clontech) and diagnostic digestion with XbaI was performed to confirm correct orientation. A hemagglutinin (HA)-tagged C->S mutant (amino acid 1095) was generated from the WT PTPµ retrovirus, pRev-TRE2-PTPµ, using a Quikchange site-directed mutagenesis kit (Stratagene, La Jolla, CA). Final sequence was confirmed by automated dideoxy DNA sequencing. The pRev-TRE2 plasmid encoding either WT PTPµ, the catalytically impaired YFDA PTPµ mutant, or the catalytically inactive C->S PTPµ mutant, each was introduced into PT67 cells (Clontech), and selected with hygromycin B 400 mg/ml (Hoffmann-La Roche Inc., Nutley, NJ). Similarly, a pRev-Tet-Off plasmid (Clontech) was transfected into PT67 cells and was selected with 800 mg/ml of G418 (Invitrogen, Carlsbad, CA). The packaged retroviral particles containing either WT PTPµ or the YFDA mutant were presented to subconfluent HMEC-1 cells, which were then selected with 200 mg/ml of hygromycin B. For experiments with the PTPµ C->S mutant, HMEC-1 cells were simultaneously co-infected with pRev-TRE-2 C->S mutant and pRev-Tet-Off. These cells were selected with both 200 mg/ml of hygromycin B and 400 mg/ml of G418 and the stably infected selectants cultured in the presence or absence of 1 µg/ml of doxycycline. Total (ectopic and endogenous) PTPµ protein expression was monitored with quantitative PTPµ immunoblotting with anti-PTPµ SK7 antibody whereas ectopic PTPµ expression was determined by immunoblotting with anti-HA antibody.

Assay for Endothelial Barrier Development

Transendothelial 14C-bovine serum albumin (BSA) flux was used as a measure of endothelial paracellular permeability as previously described.3,4 Human lung microvascular ECs were seeded at 2.5 x 105 cells in 0.5 ml of media per assay chamber and cultured to confluence. After establishment of the baseline barrier function of each monolayer, ECs were transfected with either PTPµ or control siRNAs or incubated with either the transfection reagent or media alone. After 3 hours, the monolayers were washed and transendothelial 14C-BSA flux assayed every 24 hours. In other experiments, HMEC-1 cells stably infected with pRev-TRE2 encoding for either WT PTPµ, the catalytically impaired YFDA PTPµ mutant, the catalytically inactive C->S PTPµ mutant, or the empty vector alone, each were seeded at an equivalent density of 20,000 ECs in 0.5 ml of media onto gelatin-impregnated polycarbonate filters (13-mm diameter, 0.4-µm pore size) (Nucleopore Inc.) mounted in polystyrene chemotactic chambers (ADAPS) inserted into the wells of 24-well plates. The cells were cultured for 48 hours, after which an equivalent amount of tracer molecule, 14C-BSA (specific activity, 89 µCi/mg protein; Sigma Chemical Co., St. Louis, MO), was applied to each upper compartment for 1 hour at 37°C, after which the lower compartment was counted for 14C activity. The cells stably co-infected with both pRev-TRE2 encoding for the C->S PTPµ mutant and pRev-Tet-Off were cultured in the presence and absence of 1 µg/ml of doxycycline. Transendothelial 14C-BSA flux was expressed as pmol/h.

Co-Immunoprecipitation Assays

Postconfluent human lung microvascular ECs were thoroughly rinsed with ice-cold HEPES buffer and solubilized with a low-stringency lysis buffer containing 20 mmol/L Tris, pH 7.5, 2 mmol/L CaC2, 1% Triton X-100, 5 mg/ml leupeptin, 5 mg/ml aprotinin, 1 mmol/L benzamidine, 200 µmol/L PAO, 1 mmol/L vanadate, and 0.1 mmol/L molybdate as described.29,50 The EC lysates were precleared by incubation for 1 hour at 4°C with protein G cross-linked to agarose (Sigma), preloaded with a species- and isotype-matched irrelevant antibody (AFAP IgG1, Transduction Laboratories). The lysates were then incubated overnight at 4°C with anti-PTPµ (SK7) antibody (7.5 µg of antibody/500 µg of lysate)49 or an equivalent concentration of the irrelevant antibody control. The resultant immune complexes were immobilized by incubation with protein G cross-linked to agarose for 2 hours at 4°C, centrifuged, washed five times, boiled for 7 minutes in sample buffer, and again centrifuged. The supernatants were processed for immunoblotting as described above. The blots were probed with specific murine monoclonal antibodies raised against VE-cadherin (cadherin-5, Transduction Laboratories; 30Q8A and 30Q6F, ICOS Corp., Bothell, WA). Each of the three anti-VE-cadherin antibodies (cadherin-5, 30Q8A, and 30Q6F) were raised against peptides corresponding to portions of the NH2-terminal ectodomain. In other experiments, VE-cadherin immunoprecipitates were similarly processed and probed with anti-PTPµ antibodies. The anti-VE-cadherin immunoprecipitating antibody used was 30Q8A (ICOS Corp.). The blots were subsequently incubated with HRP-conjugated anti-mouse IgG (Transduction Laboratories) and developed with ECL. To control for loading and transfer of immunoprecipitates, blots were stripped and reprobed with the immunoprecipitating antibody followed by HRP-conjugated, species-appropriate secondary IgG and were developed with ECL.

Glutathione S-Transferase (GST)-PTPµ- and VE-Cadherin-Binding Assays

EC lysates were incubated for 3 hours at 4°C with GST fusion proteins of the COOH-terminal cytoplasmic domain of either VE-cadherin (amino acids 629 to 793)28 or PTPµ (amino acids 774 to 1452)49 coupled to glutathione-Sepharose 4B beads (Pharmacia, Piscataway, NJ). The PTPµ-and VE-cadherin-binding proteins bound to the beads were extensively washed, boiled in sample buffer, resolved by SDS-polyacrylamide gel electrophoresis (PAGE), and transferred to PVDF membrane. The GST-PTPµ-binding proteins were probed with anti-VE-cadherin (30Q8A, ICOS Corp.) antibodies and the GST-VE-cadherin-binding proteins were probed with anti-PTPµ antibodies. Simultaneous GST bead controls were performed. In selected experiments, the GST-fusion proteins each containing either a thrombin or factor Xa recognition site, were subjected to protease cleavage with either thrombin (1 U/µl PBS) or factor Xa (1 U/µl in 50 mmol/L Tris-HC1, 150 mmol/L NaCl, 1 mmol/L CaCl2, pH 7.5) cleavage buffers in a GST trap column (Amersham Pharmacia, Piscataway, NJ) to remove the GST tag. After cleavage, proteins were eluted off the column, collected, resolved by SDS-PAGE, and predicted gel mobility confirmed with protein staining. Purified recombinant VE-cadherin was incubated for 3 hours at 4°C with either GST-PTPµ or GST-ß-catenin each coupled to beads or with beads alone whereas purified recombinant PTPµ was incubated with either GST-VE-cadherin or GST-ß-catenin each coupled to beads or with beads alone. The PTPµ-binding and ß-catenin-binding proteins were processed for immunoblotting with affinity-purified, goat polyclonal anti-VE-cadherin antibodies raised against a peptide corresponding to the COOH-terminal cytoplasmic domain (Santa Cruz Biotechnology Inc.) and VE-cadherin-binding and ß-catenin-binding proteins were processed for immunoblotting with anti-PTPµ antibodies. Purified recombinant proteins were used as simultaneous positive controls.

Effect of Overexpression of WT PTPµ on Tyrosine Phosphorylation State of VE-Cadherin

HMEC-1 cells were stably infected with either WT PTPµ, or the empty pRev-TRE2 vector, and cultured in the absence of doxycycline. Cells were lysed, and the lysates immunoprecipitated with either anti-VE-cadherin (30Q8A, ICOS) or anti-ß-catenin (Transduction Laboratories) antibodies as described in the co-immunoprecipitation assays above. The VE-cadherin and ß-catenin immunoprecipitates were resolved by SDS-PAGE, transferred to PVDF membrane, and the blots probed with antiphosphotyrosine antibody (PY-plus; Zymed, South San Francisco, CA) as previously described in detail.3,4 To control for loading and transfer of immunoprecipitates, blots were stripped and reprobed with the immunoprecipitating antibody followed by HRP-conjugated species appropriate secondary IgG. The blots were developed with ECL and the bands of interest normalized to the appropriate loading control.

Statistical Methods

Analysis of variance was used to compare the mean responses among experimental and control groups for all experiments. The Dunnett and Scheffé F-tests were used to determine between which group’s significant differences existed. A P value of <0.05 was considered significant.


   Results
Top
 Abstract
 Materials and Methods
 Results
Discussion
 References
 
PTPµ Expression and Immunolocalization in Pulmonary Vascular Endothelia

Human pulmonary artery and lung microvascular ECs were cultured to confluence under identical conditions. Using RT-PCR, PTPµ mRNA was detected in both endothelia (Figure 1A) . EC lysates were immunoblotted with antibodies raised against PTPµ (Figure 1B) . To ensure equal protein loading, blots were stripped and reprobed for ß-tubulin. In both endothelia, PTPµ-immunoreactive bands that migrated with apparent Mr of 200,000 and 100,000 were revealed. On a 6% gel, the 200-kd band resolved into a doublet with apparent Mr of ~210,000 and 195,000. PTPµ protein expression in the two endothelia was comparable. Therefore, PTPµ is expressed at both the mRNA and protein levels in both pulmonary artery and lung microvascular ECs where full-length PTPµ is proteolytically processed into ~100-kd cleavage products as has been described in both epithelia23,42 and HUVECs.24



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  		Figure 1. PTPµ expression and immunolocalization in pulmonary vascular endothelia. A: RT-PCR was used to detect PTPµ mRNA in postconfluent human pulmonary artery ECs (HPAEC)s and lung microvascular ECs (HPMVECs). The control DNA ladder is shown on the left and the negative control is on the right. B: PTPµ immunoblotting was used to screen for PTPµ protein in pulmonary vascular ECs. Lysates from postconfluent ECs were resolved by SDS-PAGE and were transferred to PVDF membranes. The blots were probed with murine monoclonal anti-PTPµ antibodies (SK7) followed by HRP-conjugated anti-mouse IgG and developed with ECL. To confirm equivalent protein loading, each blot was stripped and reprobed with anti-ß-tubulin antibody. Mr of protein standards in kd are shown on the right. Arrows on right indicate PTPµ-immunoreactive bands at ~200 kd and ~100 kd. Each blot is representative of two experiments. Human pulmonary artery (C) and lung microvascular (D) ECs, each were cultured to postconfluence, fixed, permeabilized, blocked, incubated with murine anti-PTPµ SK7 antibodies followed by FITC-conjugated anti-mouse IgG, and analyzed by epifluorescence microscopy. Arrows indicate PTPµ signal at intercellular boundaries. Original magnifications, x750.

 
To determine the subcellular localization of PTPµ in pulmonary vascular ECs, postconfluent pulmonary artery and lung microvascular ECs were probed with murine anti-PTPµ (SK7) antibodies followed by FITC-conjugated goat anti-mouse IgG and analyzed by epifluorescence microscopy (Figure 1, C and D) . Both endothelia displayed a fluorescence signal that was almost exclusively restricted to the intercellular boundaries, and to a much lesser degree, to perinuclear regions. These results indicate that in postconfluent pulmonary vascular ECs, PTPµ is enriched to EC-EC junctions.

PTPµ Regulates Endothelial Barrier Function

To determine whether PTPµ might influence EC-EC association and monolayer barrier function, postconfluent human lung microvascular EC monolayers were transfected with either PTPµ-targeting or control siRNA (Figure 2A) . PTPµ protein was knocked down >95% compared to the simultaneous controls from days 1 to 5. PTPµ protein abundance in ECs transfected with control siRNA was no different from that seen in the media control (Figure 2A , lane 1 versus lanes 2, 4, 6, 8, 10, and 12). 14C-BSA flux across monolayers transfected with PTPµ siRNA was dramatically increased on days 3, 4, 5, and 6 compared to monolayers transfected with the control siRNA (P < 0.02) (Figure 2B) . Throughout this same time period, mean 14C-BSA flux across monolayers transfected with control siRNA was not different from flux across the simultaneous media controls. On days 1 and 2, 14C-BSA flux across monolayers transfected with either control or PTPµ siRNA was increased compared to the simultaneous media controls but was not significantly different from each other or from flux across monolayers incubated with the transfection reagent alone (data not shown). These data indicate that the early barrier dysfunction could be ascribed to the transfection reagent whereas the loss of barrier function on day ≥3 was because of selective knockdown of PTPµ.



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  		Figure 2. Effect of knockdown of PTPµ through siRNA transfection on endothelial barrier function. A: Postconfluent human lung microvascular ECs were transfected with PTPµ-targeting (lanes 3, 5, 7, 9, 11, and 13) or control (lanes 2, 4, 6, 8, 10, and 12) siRNA and after increasing times (days 1 to 6), were lysed and the lysates resolved by SDS-PAGE, transferred to PVDF membrane, and immunoblotted for PTPµ. To confirm equivalent loading, each blot was stripped and reprobed with anti-ß-tubulin antibody. B: Human lung microvascular ECs were cultured to confluence in barrier function assay chambers when baseline barrier function was established (day 0). The postconfluent ECs were cultured in media alone or were transfected with PTPµ-targeting or control siRNAs after which transendothelial 14C-BSA flux was assayed every 24 hours. Vertical bars represent mean (±SE) 14C-BSA flux in pmol/hour across media control EC monolayers or monolayers transfected with either PTPµ or control siRNA. *, Significantly increased compared to the simultaneous media control monolayers at P < 0.02. **, Significantly increased compared to the simultaneous siRNA control at P < 0.006.

 
In selected experiments, HMEC-1 cells, immortalized human dermal microvascular ECs, were used to manipulate PTPµ expression and catalytic activity. To help define this experimental system, we determined whether HMEC-1 cells express PTPµ and ZA proteins and whether their paracellular pathway was responsive to PTP inhibition.28 Using RT-PCR, PTPµ mRNA was confirmed in HMEC-1 cells (Figure 3A) . Lysates from postconfluent HMEC-1 cells were processed for immunoblotting for PTPµ (Figure 3B) , as well as VE-cadherin, and {alpha}-, ß-, {gamma}-, and p120-catenins (Figure 3C) . PTPµ-immunoreactive bands that migrated with apparent Mr of 200,000 and 100,000 were revealed. Of note, the ~200-kd band did not migrate as a doublet as seen in the two primary cultured pulmonary vascular endothelia (Figure 1B) . VE-cadherin and its associated catenins each were expressed with the anticipated gel mobilities (Figure 3C) . PTP inhibition with vanadate resulted in concentration-dependent loss of barrier function in postconfluent HMEC-1 monolayers (Figure 3D) . Therefore, HMEC-1 cells express PTPµ and all five ZA proteins for which we screened and contains a paracellular pathway that responds to PTP inhibition.



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  		Figure 3. PTPµ and ZA protein expression and barrier responsiveness to PTP inhibition in HMEC-1 cells. A: RT-PCR was used to detect PTPµ mRNA in HMEC-1 cells. The control DNA ladder is shown on the left and the negative control is on the right. B and C: Lysates of postconfluent HMEC-1 cells were resolved by SDS-PAGE, transferred to PVDF membrane, and immunoblotted for PTPµ (B), VE-cadherin, and {alpha}-, ß-, {gamma}-, and p120 catenins (C). C: The blot was stripped and reprobed for ß-tubulin. Molecular weight markers in kd are indicated. D: Vertical bars represent mean (±SE) transendothelial 14C-BSA flux in pmol/hour immediately after 6 hours of exposure to increasing concentrations of vanadate or media alone. n indicates the number of monolayers studied. Mean (±SE) 14C-BSA flux across naked filters without EC monolayers is shown. *, Significantly increased compared to the simultaneous media control monolayers at P < 0.05.

 
HMEC-1 cells overexpressing WT PTPµ, the catalytically impaired PTPµ YFDA mutant, or the empty vector were selected with hygromycin B. HMEC-1 cell lysates from pooled selectants were resolved by SDS-PAGE, transferred to PVDF membrane, and the blots probed with anti-PTPµ SK7 antibody. Full-length (~200 kd) PTPµ expression in HMEC-1 cells stably infected with either WT PTPµ or the YFDA mutant was increased ~7- to 20-fold relative to cells infected with the empty vector (Figure 4A , inset). When ECs overexpressing either WT PTPµ or the YFDA mutant were seeded into the wells of 12-well plates (5 x 104 ECs/well), cultured for 72 hours, and stained with Trypan blue, mean (±SE) percent viability (97.24 ± 0.36, n = 3, and 94.45 ± 0.45, n = 3, respectively) was comparable to the vector control (96.53 ± 0.67%, n = 3) or parental HMEC-1 cells (96.51 ± 0.78%, n = 3).



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  		Figure 4. Effect of overexpression of WT and catalytically impaired PTPµ mutants on endothelial barrier function. A: HMEC-1 cells stably infected with pRev-TRE-2 encoding for WT PTPµ, the catalytically impaired YFDA mutant, or the empty vector control (VC), each were seeded at equivalent densities in assay chambers and transendothelial 14C-BSA flux assayed at 48 hours. *, Significantly decreased compared to the vector control at P < 0.05. **, Significantly increased compared to the vector control at P < 0.05. Inset: PTPµ immunoblot of lysates of pooled selectants of HMEC-1 cells stably infected to overexpress WT PTPµ, the vector control, or the YFDA mutant. B: HMEC-1 cells stably co-infected with pRev-TRE-2 encoding for the catalytically inactive C->S PTPµ mutant and pRev-Tet-Off were cultured for 48 hours in assay chambers in the presence and absence of doxycycline after which transendothelial 14C-BSA flux was assayed. *, Significantly increased compared to the doxy + control at P < 0.002. Inset: HA immunoblot of lysates of pooled selectants of HMEC-1 cells stably co-infected to overexpress the HA-tagged C->S PTPµ mutant after withdrawal of doxycycline. Arrows on left indicate 200-kd full-length and 100-kd proteolytically cleaved PTPµ protein. In both A and B, each vertical bar represents mean (±SE) 14C-BSA flux in pmol/hour across EC monolayers overexpressing WT PTPµ, the vector control, the catalytically impaired YFDA mutant, or the tetracycline-responsive, catalytically inactive C->S mutant. n is indicated in each bar.

 
HMEC-1 cells overexpressing WT PTPµ or the empty vector alone were cultured in barrier function assay chambers for 48 hours after which transendothelial 14C-BSA flux was assayed (Figure 4A) . The vector control cells achieved tight barrier function. The pooled and selected HMEC-1 cells overexpressing WT PTPµ formed monolayers that permitted even less transendothelial albumin flux than did the vector control (P < 0.05). In contrast, comparable overexpression of the catalytically impaired YFDA PTPµ mutant failed to enhance barrier function (Figure 4A) . In other experiments, HMEC-1 cells stably co-infected with pRev-TRE2 encoding for the catalytically inactive C->S PTPµ mutant and pRev-Tet-Off were cultured in barrier function assay chambers in the presence and absence of 1 µg/ml of doxycycline for 48 hours after which transendothelial 14C-BSA flux was assayed (Figure 4B) . After removal of doxycycline, both ectopic expression of the C->S PTPµ mutant and 14C-BSA flux were increased (P < 0.002) compared to expression and flux in the presence of doxycycline. Although increased expression of WT PTPµ decreased albumin flux, increased expression of either of two catalytically impaired PTPµ mutants, each with only one (C->S) or two (YFDA) amino acid substitutions, failed to do so (Figure 4, A and B) . These combined data indicate that full catalytic activity is required for PTPµ to enhance barrier function. That overexpression of either catalytically impaired mutant failed to enhance barrier function indicates that overexpression of intact PTPµ ectodomain, together with its regulatory noncatalytic domains, was insufficient to reproduce the WT PTPµ effect. Overexpression of either of the two catalytically impaired PTPµ mutants not only failed to display the barrier-enhancing effect of WT PTPµ, each profoundly impaired barrier function (Figure 4, A and B) .

PTPµ Interacts with VE-Cadherin

Since PTPµ could be localized to the intercellular boundaries of postconfluent ECs, we asked whether it associated with the membrane-spanning, EC-restricted cadherin, VE-cadherin. Immunoprecipitation of PTPµ with SK7 in lung microvascular ECs lysates, co-immunoprecipitated VE-cadherin (Figure 5A) compared to the simultaneous bead controls preloaded with a species- and isotype-matched irrelevant antibody. In reciprocal manner, immunoprecipitation of VE-cadherin co-immunoprecipitated both full-length PTPµ and the ~100-kd PTPµ fragments (Figure 5B) . In other experiments, lysates of postconfluent human lung microvascular ECs were incubated with GST fusion proteins of either the intracellular segment of PTPµ (Figure 5C) , or the cytoplasmic domain of human VE-cadherin (Figure 5D) , each coupled to glutathione Sepharose beads, or incubated with a GST bead control. The PTPµ-binding proteins were processed for immunoblotting with either of two distinct anti-VE-cadherin antibodies (Figure 5C) . The GST-PTPµ-bound VE-cadherin was detected by either of the two anti-VE-cadherin antibodies whereas none was detected in the GST bead control. When the VE-cadherin-binding proteins were probed for PTPµ, GST-VE-cadherin bound PTPµ compared to the GST bead control (Figure 5D) . Each of the two PTPµ-immunoreactive bands in the doublet bound to GST-VE-cadherin. These data indicate that PTPµ directly or indirectly interacts with VE-cadherin in vitro. To determine whether PTPµ co-localizes with VE-cadherin in an intact EC system, immunofluorescence microscopy was applied using simultaneous dual-antibody labeling (Figure 6) . As anticipated, PTPµ (Figure 6C) and VE-cadherin (Figure 6A) each localized to intercellular boundaries. Merging the images obtained from identical fields with antibodies against each of these proteins revealed a high degree of co-localization of PTPµ and VE-cadherin in intercellular junctions (Figure 6B , regions of co-localization appear yellow). To determine whether the PTPµ-VE-cadherin interaction might be direct, the in vitro binding assays were performed with purified recombinant proteins in the absence of EC lysates (Figure 7) . Purified recombinant VE-cadherin (amino acids 629 to 723) cleaved from GST was incubated with GST-PTPµ coupled to beads or with beads alone (Figure 7A) . In reciprocal pull-down experiments, purified recombinant PTPµ (amino acids 774 to 1452) cleaved from GST was incubated with GST-VE-cadherin coupled to beads or beads alone (Figure 7B) . The purified recombinant VE-cadherin bound to GST-PTPµ as well as the GST-ß-catenin-positive control but not to the bead control (Figure 7A) and the purified recombinant PTPµ bound to the GST-VE-cadherin but not to either GST-ß-catenin or the bead control (Figure 7B) . These combined data indicate that PTPµ directly and specifically associates with VE-cadherin.



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  		Figure 5. PTPµ-VE-cadherin interactions. A and B: PTPµ and VE-cadherin co-immunoprecipitation assays. Lysates of postconfluent lung microvascular ECs were immunoprecipitated with either anti-PTPµ (SK7) or anti-VE-cadherin (30Q8A, ICOS) antibodies. The immunoprecipitates were resolved by SDS-PAGE, transferred onto PVDF membrane, and the blots incubated with either anti-VE-cadherin (A) or anti-PTPµ (B) antibodies followed by HRP-conjugated anti-murine antibody, and developed with ECL. Arrows on right indicate 200-kd and 100-kd PTPµ-immunoreactive bands. In A, the positive control is a total cell lysate indicated by —. C and D:GST-PTPµ- and GST-VE-cadherin-binding assays. Lysates from postconfluent lung microvascular ECs were incubated for 3 hours at 4°C with GST fusion proteins of the cytoplasmic domains of either PTPµ (C) or VE-cadherin (D) coupled to beads or incubated with beads alone. The PTPµ- and VE-cadherin-binding proteins were washed, boiled in sample buffer, resolved by SDS-PAGE, and transferred to PVDF membrane. The blots of PTPµ-binding and VE-cadherin-binding proteins were probed with two distinct anti-VE-cadherin antibodies (30Q8A, ICOS, and cadherin-5; Transduction Laboratories) and an anti-PTPµ antibody, respectively, followed by HRP-conjugated anti-murine antibody and developed with ECL. Whole EC lysates without previous incubation with GST-PTPµ or GST-VE-cadherin were immunoblotted as positive controls that are indicated by —. The molecular weights in kd are indicated on the left. Arrows on right indicate 200-kd and 100-kd PTPµ-immunoreactive bands. IP, immunoprecipitate; IB, immunoblot; bc, bead control. Each blot is representative of three experiments.

 


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  		Figure 6. Co-localization of PTPµ and VE-cadherin. Postconfluent human lung microvascular ECs were cultured on fibronectin (25 mg/ml)-coated glass coverslips for 48 hours, after which they were fixed and permeabilized with 3% paraformaldehyde and 0.5% Triton X-100, and stained with goat polyclonal anti-VE-cadherin (Santa Cruz) and monoclonal SK7 anti-PTPµ antibodies, followed by fluorescein- and Cy-3-labeled secondary antibodies, respectively. Nuclei were visualized with bis-benzimide. A: VE-cadherin. B: Merged image of VE-cadherin and PTPµ with Hoescht reagent. Regions of co-localized VE-cadherin and PTPµ appear yellow and are indicated by the arrows and nuclei appear blue. C: PTPµ. Scale bar, 20 µm. Original magnifications, x600.

 


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  		Figure 7. GST-PTPµ and GST-VE-cadherin-binding assays with purified recombinant binding partners. The GST-PTPµ- and GST-VE-cadherin-binding assays were repeated as described in Figure 5, C and D , but in these experiments the GST fusion proteins coupled to beads were incubated with recombinant binding partners that were proteolytically cleaved from GST and purified in a GST trap column. GST-PTPµ and GST-ß-catenin were incubated with purified VE-cadherin cleaved from GST (A) and GST-VE-cadherin and GST-ß-catenin were incubated with purified PTPµ cleaved from GST (B). In each case, simultaneous empty bead controls were performed. The PTPµ-binding, VE-cadherin-binding, and ß-catenin-binding proteins were processed for immunoblotting with goat anti-human VE-cadherin (A) (Santa Cruz) and anti-PTPµ (SK7) (B) antibodies, respectively. Purified recombinant proteins were used for the positive controls. The molecular weights in kd are indicated on the left. IB, immunoblot; bc, bead control. Each blot is representative of three experiments.

 
Overexpression of WT PTPµ in HMEC-1 Cells Modulates the Tyrosine Phosphorylation of VE-Cadherin

To determine whether VE-cadherin might be an in vivo substrate for PTPµ, WT PTPµ was overexpressed (Figure 8A) and changes in VE-cadherin and ß-catenin tyrosine phosphorylation sought (Figure 8B) . Lysates of HMEC-1 cells overexpressing WT PTPµ or the empty vector alone were immunoprecipitated with antibodies raised against VE-cadherin or ß-catenin and the immunoprecipitates processed for phosphotyrosine immunoblotting (Figure 8B) . In ECs overexpressing WT PTPµ, tyrosine phosphorylation of VE-cadherin (Figure 8B , lanes 1 and 2) was decreased compared to the vector controls, suggesting that VE-cadherin is a potential in vivo substrate for PTPµ catalytic activity. In contrast, overexpression of WT PTPµ did not decrease tyrosine phosphorylation of ß-catenin (Figure 8B , lanes 3 and 4). Therefore, overexpression of PTPµ selectively dephosphorylated VE-cadherin without modifying another closely associated ZA component, ß-catenin. It is conceivable that other as yet unidentified PTPµ substrates are also involved. Phosphotyrosine immunoblotting of total cell lysates of ECs, in which either WT PTPµ was overexpressed ~10-fold or >90% PTPµ was knocked down with PTPµ siRNA, failed to demonstrate any consistent differences in phosphotyrosine signal compared to their respective controls (data not presented).



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  		Figure 8. Overexpression of WT PTPµ modulates tyrosine phosphorylation states of VE-cadherin. A: HMEC-1 cells were stably infected with packaged retroviral particles containing pRev-TRE2 encoding for WT PTPµ or the empty vector control (VC), and were selected with hygromycin B. HMEC-1 cell lysates from pooled selectants were resolved by SDS-PAGE, transferred to PVDF membrane, and the blots probed with anti-PTPµ SK7 antibody followed by HRP-conjugated anti-murine antibody and developed with ECL. The arrows on the right indicate the ~200-kd full-length PTPµ and the ~100-kd cleavage products. B: Lysates of HMEC-1 cells stably infected with pRev-TRE2 encoding for WT PTPµ or the empty vector (VC), each were immunoprecipitated with antibodies raised against VE-cadherin (30Q8A, ICOS) and ß-catenin. In B, the VE-cadherin and ß-catenin immunoprecipitates were resolved by SDS-PAGE, transferred to PVDF membrane, and the blots incubated with HRP-conjugated antiphosphotyrosine PY-plus antibody and developed with ECL. For normalization of the phosphotyrosine signal to the immunoprecipitated protein, blots were stripped and reprobed with the immunoprecipitating antibodies. IP, immunoprecipitate; IB, immunoblot; IB*, immunoblot after stripping; VC, vector control. Molecular weights in kd are indicated on each side. Each blot is representative of more than two experiments.

 

   Discussion
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 Abstract
 Materials and Methods
 Results
 Discussion
References
 
In this report, we have demonstrated that PTPµ is expressed at the mRNA and protein levels in both human pulmonary artery and lung microvascular endothelia where it is enriched to EC-EC junctions. In both endothelia, PTPµ-immunoreactive bands that migrated with apparent Mr of 200,000 and 100,000 were revealed and protein expression in the two endothelia were comparable. Overexpression of WT PTPµ enhanced endothelial barrier function whereas comparable overexpression of either of two catalytically impaired PTPµ mutants not only failed to tighten the barrier but profoundly diminished it. These results indicate that an intact catalytic domain is required for the barrier-enhancing activity of WT PTPµ. Whereas overexpression of WT PTPµ enhanced barrier function, knockdown of PTPµ through siRNA opened the paracellular pathway. In postconfluent human lung microvascular ECs, the cytoplasmic domain of PTPµ directly associated with the intracellular segment of VE-cadherin. Overexpression of WT PTPµ in HMEC-1 cells demonstrated VE-cadherin as a potential in vivo PTPµ substrate. These combined data suggest that PTPµ is expressed in pulmonary vascular endothelia where it directly associates with and regulates the tyrosine phosphorylation state of VE-cadherin as well as EC-EC barrier integrity.

A tyrosine phosphorylation-responsive endothelial paracellular pathway, in which agonists that increase paracellular permeability also increase tyrosine phosphorylation of one or more ZA proteins, has been described.3-6 PTPs are expressed in various epithelia where they associate with and/or dephosphorylate ZA proteins.29-34 However, less is known about ZA-associated PTPs in endothelia.7,35,36,42 We asked whether PTPµ may provide such counterregulation within the human pulmonary vascular endothelium. Recently, PTPµ protein was found to be expressed in vascular ECs in adult rats and swine although PTPµ immunostaining was more prominent in arteries, arterioles, and the vasa vasorum than in veins and capillaries.42 In the current report, we found that in human postconfluent pulmonary artery and lung microvascular ECs, PTPµ protein expression in the two lung endothelia in vitro was comparable (Figure 1, A and B) . PTPµ expression in EC monolayers reportedly increases with confluence.24 It is conceivable that in cultured, contact-inhibited ECs, PTPµ expression approaches a maximal level whereas in vivo, other environmental stimuli are operative.41 In both pulmonary vascular endothelia, full-length PTPµ resolved into a doublet (Figure 1B) . Although in most reports full-length PTPµ appears as a single band, the PTPµ doublet has been described in COS cells transiently transfected with full-length PTPµ,24 Sf9 insect cells infected with recombinant baculovirus expressing full-length PTPµ,38 and in disassociated cells in chick retinal explants.51 Whether the ~200-kd doublet in our EC system represents alternatively spliced variants, multiple phosphorylation states, proteolytic processing, and/or other posttranslational modifications is unclear. Such alternative splicing events have been described in the closely related receptor PTP, PTP{rho}.52

To evaluate whether PTPµ may regulate endothelial barrier function, PTPµ protein expression in ECs was genetically manipulated. In human lung microvascular ECs, PTPµ depletion through RNA interference disrupted barrier function (Figure 2) indicating that PTPµ, directly or indirectly, is absolutely required for barrier maintenance. In HMEC-1 cells, overexpression of WT PTPµ enhanced barrier function (Figure 4A) . In contrast, comparable overexpression of either of two catalytically impaired PTPµ mutants, each with only one or two amino acid substitutions, displayed no barrier-enhancing activity (Figure 4A) . These findings suggest that the ability of WT PTPµ to promote barrier function requires an intact catalytic domain and may be mediated through tyrosine dephosphorylation of VE-cadherin. Overexpression of the C->S PTPµ mutant that contains an intact PTPµ ectodomain but no catalytic activity did not diminish transendothelial albumin flux (Figure 4, A and B) . This indicates that increased homophilically interacting ectodomain alone does not enhance paracellular pathway function. Similarly, overexpression of the putative regulatory, noncatalytic domains of PTPµ was insufficient to enhance barrier function. That increased availability of these potential, protein-binding domains does not tighten barrier function suggests that their increased association with potential binding partners does not explain the barrier-enhancing activity. Not only did each of the two catalytically impaired PTPµ mutants fail to simulate the WT PTPµ barrier-enhancing effect, they dramatically compromised barrier function (Figure 4, A and B) . Overexpression of either of these two catalytically impaired, substrate-trapping mutants may sequester binding partners competitively displacing WT PTPµ thereby preventing downstream signaling events.

Since PTPµ could be localized to the intercellular boundaries of postconfluent lung microvascular ECs (Figures 1C and 6C) , we asked whether it associated with the established EC-EC adherens junctional protein VE-cadherin. In the extracts of rat lung and various cell lines, PTPµ interacts with the classical cadherins, E-cadherin, N-cadherin, and cadherin-4.30,53 Although multiple cadherins can be co-expressed and differentially distributed in ECs,12 VE-cadherin is unique to ECs and is localized to their intercellular junctions where they regulate paracellular pathway function.13 Among the classical cadherins, the cytoplasmic domain of VE-cadherin is the least conserved.48 It shares only ~36% amino acid identity with E-cadherin. It was therefore unclear whether PTPµ also binds VE-cadherin. In recent double-label co-localization fluorescent microscopy studies in HUVECs, PTPµ co-localized with VE-cadherin.42 We now have demonstrated in a cell-free system, that purified recombinant cytoplasmic domain of PTPµ binds to immobilized GST-VE-cadherin (Figure 7B) and purified recombinant cytoplasmic domain of VE-cadherin binds to immobilized GST-PTPµ (Figure 7A) . These data clearly indicate that the cytoplasmic domain of PTPµ directly associates with the intracellular segment of VE-cadherin. The COOH-terminal 38-amino acid portion of E-cadherin is required for its association with PTPµ.29,30 The same sequence in human VE-cadherin shares ~60% identity and ~80% similarity with E-cadherin (BLAST, www.ncbi.nih.gov). It may be through this PTPµ-VE-cadherin interaction that PTPµ localizes to the EC-EC adherens junction.

To determine whether VE-cadherin could be an in vivo PTPµ substrate, we manipulated PTPµ catalytic activity in HMEC-1 cells. Overexpression of full-length WT PTPµ, containing the two catalytic domains and potential protein-binding domains, consistently resulted in tyrosine dephosphorylation VE-cadherin (Figure 8B) . If the intracellular pool of a PTPµ-binding partner is finite or rate limiting, overexpression of WT PTPµ might indiscriminately dephosphorylate substrates that are in close proximity. In the current studies, a wide range of levels of WT PTPµ overexpression consistently demonstrated VE-cadherin as a substrate for tyrosine dephosphorylation (data not shown) whereas the VE-cadherin-binding partner, ß-catenin, remained unchanged (Figure 8B) . It is possible that other as yet unidentified PTPµ substrates are also involved. It is also possible that PTPµ regulates the paracellular pathway independent of its catalytic activity. Of note, introduction of PTPµ in prostate carcinoma cells that lack PTPµ restores E-cadherin-dependent adhesion but does so independent of PTPµ catalytic activity.54

VE-cadherin appears to be a potential in vivo substrate for PTPµ (Figure 8B) . Although VE-cadherin has been implicated in multiple aspects of EC behavior,55 it is unclear whether any of these VE-cadherin functions or VE-cadherin-protein interactions are tyrosine phosphorylation-dependent. In one study, vascular endothelial growth factor-induced tyrosine phosphorylation of VE-cadherin in HUVECs was temporally coincident with increases in both paracellular permeability to FITC-dextran and EC migration.6 In another study, tumor necrosis factor-{alpha} similarly increased both tyrosine phosphorylation of VE-cadherin and movement of a permeability tracer across EC monolayers.20 These studies imply that increased VE-cadherin tyrosine phosphorylation is associated with opening of the endothelial paracellular pathway. Whether these and other tumor necrosis factor-{alpha}- and vascular endothelial growth factor-induced EC responses, including angiogenesis, can be causally related to VE-cadherin tyrosine phosphorylation, and whether PTPµ might counterregulate one or more of these VE-cadherin functions is unknown. In the current studies, overexpression of WT PTPµ decreased tyrosine phosphorylation of VE-cadherin (Figure 8B , lanes 1 and 3) and enhanced barrier function (Figure 4) . These combined data indicate that PTPµ regulates both the tyrosine phosphorylation state of VE-cadherin and paracellular pathway function. Whether a causal relationship between these two EC responses exists and the mechanism(s) through which the intracellular modification might be coupled to functional changes outside the cell remain unknown. Interestingly, there is evidence that PTPµ regulates N-cadherin function.44 Down-regulation of PTPµ catalytic activity through either anti-sense or overexpression of a catalytically inactive PTPµ mutant decreased neurite outgrowth on an N-cadherin substrate, suggesting that PTPµ regulates N-cadherin-mediated homophilic adhesion.

The regulation of EC-EC interactions and the paracellular pathway through protein tyrosine phosphorylation is not well understood. Our findings indicate that in pulmonary vascular ECs, PTPµ directly associates with the cytoplasmic domain of VE-cadherin. That PTPµ can directly interact with the cadherin-catenin complex suggests its importance to ZA organization and function. Here, PTPµ dephosphorylates VE-cadherin and possibly other substrates. Such modification of VE-cadherin may, through inside-to-outside signaling, regulate VE-cadherin ectodomain homophilic adhesion. Under physiological conditions, PTPµ, and possibly other PTPs, appears to maintain basal endothelial barrier function through the restraint of tyrosine phosphorylation within the ZA multiprotein complex. Loss of this counterregulatory dephosphorylation may be operative during pathological conditions associated with opening of the pulmonary microvascular endothelial paracellular pathway (eg, acute respiratory distress syndrome) and dysregulated angiogenesis (eg, diabetic retinopathy). The mechanism(s) through which PTPµ regulates EC-EC engagement/disengagement may have implications for tissue morphogenesis and remodeling, and angiogenesis within the context of wound healing and tumor cell survival.


   Acknowledgements
 
We thank Ms. Shirley A. Taylor for excellent secretarial support and Mr. Seth Crawford for photography and graphics.


   Footnotes
 
Address reprint requests to Simeon E. Goldblum, M.D., Mucosal Biology Research Center, University of Maryland School of Medicine, 22 Penn St., HSF II, Room 351, Baltimore, MD 21201. E-mail: sgoldblu{at}mbrc.umaryland.edu

Supported by the National Institutes of Health (grants HL63217, HL70155, and HL58064 to S.E.G. and GM55989 to N.K.T.) and the American Heart Association (grant 0151465U to S.E.G.).

This article is featured in a commentary by A. Verin (Am J Pathol 2005, 166:955–957), published in this issue.

Accepted for publication November 4, 2004.


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