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Cardiovascular Research Institute, Departments of Surgery and Medical Physiology, Texas A&M University System Health Science Center, 702 Southwest H. K. Dodgen Loop, Temple, TX 76504, USA
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(Received 10 July 2003;
accepted after revision 20 October 2003;
first published online 24 October 2003)
Corresponding author S. Y. Yuan: Cardiovascular Research Institute, Department of Surgery, Texas A&M University System Health Science Center, 702 Southwest H. K. Dodgen Loop, Temple, TX 76504, USA. Email: yuan{at}tamu.edu
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Introduction |
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The adhesion between endothelial cells through AJs is mainly mediated by calcium-dependent homophilic binding of the VE-cadherin molecules. VE-cadherin consists of an extracellular domain with five homologous calcium-binding repeats (EC1–EC5), a short transmembrane region, and a cytoplasmic domain (Yap et al. 1997; Bazzoni & Dejana, 2001; Dejana et al. 2001). The extracellular domains bind to each other, forming lateral homodimers or hexamers (Yap et al. 1997; Bazzoni & Dejana, 2001; Bibert et al. 2002). The highly conserved cytoplasmic domains are anchored to the actin cytoskeleton via a family of linking proteins, catenins (Yap et al. 1997; Bazzoni & Dejana, 2001; Dejana et al. 2001). In particular, the C-terminus (82 amino acids) of VE-cadherin binds to ß-catenin, which in turn interacts with 
-catenin and other junctional proteins to form a complex that anchors to the cytoskeleton (Yap et al. 1997; Bazzoni & Dejana, 2001; Dejana et al. 2001). Recent experiments suggest that the homophilic binding of the VE-cadherin extracellular domains contributes to the initial cell-cell adhesion during culture, whereas the intracellular interaction between VE-cadherin and cytoskeletal proteins is required for the full strength of junctional adhesiveness (Navarro et al. 1995; Caveda et al. 1996; Carmeliet et al. 1999; Hordijk et al. 1999; Bazzoni & Dejana, 2001; Dejana et al. 2001). However, the functional importance of the intracellular composition and activity of VE-cadherin in the physiological regulation of vascular endothelial barrier function remains unrevealed.
The aim of this study was to determine whether interruption of the structural linkage between VE-cadherin and the cytoskeleton altered the endothelial AJ integrity and microvascular barrier function. We expressed and purified a fusion protein encompassing the VE-cadherin cytoplasmic domain (rVE-cad CPD) and used it as a molecular tool to specifically block the binding of endogenous VE-cadherin to ß-catenin. Endothelial uptake of rVE-cad CPD resulted in disorganization of AJs coupled with a significant increase in the permeability of both the endothelial monolayers and venules. The results indicated that the intracellular composition and interaction of VE-cadherin with the cytoskeleton are essential to the maintenance of vascular endothelial barrier function.
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Methods |
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Based on the published sequence of human VE-cadherin, two oligonucleotide primers, VE-cadF1 (5'-1876CGGCGGCGGCTCCGGAA1892-3') and VE-cadR (5'-2371GACCTCGGCCGCCTAATACAG2351-3'), were selected for amplification of the cytoplasmic domain cDNA. RNA samples isolated from human umbilical vein endothelial cells (HUVEC) were subjected to RT-PCR and the product was cloned into the QIAexpress pQE-30 UA vector (Qiagen). Both the PCR product and plasmid DNA of the expression construct pQE30/VE-cad-CPD were sequenced for verification of sequence and orientation of the insert. The VE-cad CPD gene was fused to the 5'-terminal 6 x His-tag coding sequence in an ORF under the control of phage T5 promoter (Fig. 1). For rVE-cad CPD expression, fresh culture of E. coli harbouring the plasmid pQE30/VE-cad-CPD was incubated with Luria-Bertani (LB) broth containing ampicillin at 37°C for 3 h and then mixed with 1 µM isopropyl-ß-D-thioglactoside (IPTG). After IPTG induction for 4 h at 37°C, the culture was harvested and the cell pellet was resuspended in B-PER reagent (Pierce) for lysis. The clarified supernatant was loaded to a prebalanced Ni-NTA (nickel-nitrilotriacetic acid) spin column (Qiagen) in which the Ni-NTA silica matrices selectively bound 6 x His-tagged proteins. After washing the columns, rVE-cad CPD was eluted in a buffer containing 250 mM imidazole, which was removed by dialysis against 20 mM Tris-HCl-buffered saline at pH 7.5. The E-TOXATE assay (Sigma) was performed for endotoxin detection of the rVE-cad CPD preparations according to the supplier's instructions.
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The rVE-cad CPD protein immobilized on Ni-NTA agarose beads was incubated at 4°C for 4 h with lysate of human umbilical vein endothelial cells (HUVEC). The beads were washed with Tris-HCl buffer containing 0.05% Triton X-100, boiled in a sample buffer, and then subjected to immunoblotting. The membrane was first blotted with a monoclonal antibody to VE-cadherin (clone F8, Santa Cruz) for detection of rVE-cad CPD. After stripping, the membrane was re-probed with a horseradish peroxidase (HRP)-labelled antibody to ß-catenin (BD Biosciences) for ß-catenin detection.
Endothelial cell culture
HUVEC (passage 1) were purchased from Clonetics (San Diego, CA, USA). Cells were routinely maintained in gelatin-coated dishes containing EGM-2 culture medium (Clonetics) supplemented with 5% fetal bovine serum, 10 ng ml-1 human epidermal growth factor, 0.4 µg ml-1 hydrocortisone, vascular endothelial growth factor, human fibroblast growth factor, ascorbic acid, heparin, 50 µg ml-1 gentamicin, and 50 ng ml-1 amphotericin B. Confluent cells were digested with 0.05% trypsin–0.02% EDTA, resuspended in growth medium and seeded to 60 mm dishes for immunoprecipitation/Western blotting or on coverslips for immunofluorescence staining. Early passages (4–7) of HUVEC were used.
Immunoprecipitation and Western blotting
HUVEC were lysed in a Tris-HCl lysis buffer containing 1% Triton X-100 and protease inhibitors. The lysate was clarified by centrifugation at 13 000 g for 15 min. Protein concentrations of the soluble fraction were measured using the Bradford's method. The insoluble pellet was resuspended in lysis buffer containing 2% SDS and boiled for 10 min. Protein concentrations in the insoluble extracts were determined by the bicinchoninic acid method. Both the soluble and insoluble fractions were separated by SDS-PAGE on 4–12% Tris-glycine gels, transferred to nitrocellulose membranes, and blotted with a monoclonal antibody to either ß-catenin or VE-cadherin. Following incubation with a secondary antibody conjugated to horseradish peroxidase, immunoreactive bands were detected using the LumiGlo chemiluminescent substrate (Cell Signaling), scanned by reflectance scanning densitometry, and quantified using the NIH Image software. For immunoprecipitation, cell lysates were incubated with protein G PLUS-agarose beads preabsorbed with an anti-ß-catenin antibody (Santa Cruz) at 4°C for 2 h. After washing and boiling, samples were fractionated with SDS-PAGE on 4–12% gels and subjected to immunoblotting as described above.
Immunocytochemistry
HUVEC grown on gelatin-coated coverslips were incubated with a transfection mixture containing rVE-cad CPD for 2 h and then fixed with 2% paraformadehyde and permeabilized with 0.2% Triton X-100. For the labelling of rVE-cad CPD, cells were incubated with an antihistidine antibody (Qiagen) for 1 h followed by incubation with fluorescein isothiocyanate (FITC)-labelled antimouse IgG (Santa Cruz). Coverslips were then mounted on slides for fluorescence microscope observation. For the determination of the endogenously expressed VE-cadherin distribution, cells were incubated with a monoclonal antibody to the VE-cadherin extracellular domain (BD Bioscience) and followed by a FITC-labelled antimouse conjugate.
In vitro permeability assay
HUVEC were seeded at 105 cells (cm2)-1 on gelatin-coated Costar Transwell membranes (VWR) and grown to confluence. Fluorescently labelled bovine serum albumin was added to the top (luminal) chamber at 1.0 mg ml-1. Samples were collected from both the luminal and abluminal (bottom) chambers and analysed with a fluorescence microplate reader. Sample readings were converted with a standard curve to albumin concentration. The permeability coefficient of albumin was determined based on the equation Pa=[Ab]/tx 1/AxV/[Lu], where [Ab] is the abluminal concentration of FITC-albumin, t is time in seconds for FITC-albumin incubation, A is area of membrane in cm2, V is the volume of the abluminal chamber, and [Lu] is the luminal concentration of FITC-albumin (Tinsley et al. 1999; Yuan et al. 2002). Control experiments were performed to measure tracer flux across gelatin-coated microporous filters without cells. Monolayers were discarded if they failed to form an effective barrier as indicated by a >20-fold decrease in Pa compared with that of gelatin-coated membranes without cells.
Measurement of venular permeability
The animals used in this study were housed and handled in accordance with the protocols approved by the Institutional Animal Care and Use Committee in compliance with the NIH guideline for experimental animal usage. Pigs weighing 9–13 kg were anaesthetized with sodium pentobarbital (30 mg kg-1, I.V.) and heparinized (250 units kg-1, I.V.). A left thoracotomy was performed and the heart was electrically fibrillated, excised and placed in 4°C physiological saline. The technique of isolation and cannulation of coronary venules has been described in detail in our previous publications (Yuan et al. 1993; Yuan & Chilian, 1995, 1997). Briefly, a venule 20–50 µm in diameter was dissected and cannulated with a micropipette on each end with a third smaller pipette inserted into the inflow micropipette. Each micropipette was connected to a reservoir to allow independent control of intraluminal perfusion pressure and flow. The vessel was interchangeably perfused with either physiological salt solution or the same perfusate containing fluorescently labelled albumin. The permeability of the vessel was quantified by measuring the ratio of transvascular flux to transmural concentration difference of the tracer (Yuan et al. 1993, 1997; Yuan & Chilian, 1995). The apparent solute permeability coefficient of albumin (Pa) was calculated using the equation Pa= (1/
If)(dIf/dt)o(r/2), where If is the initial step increase in fluorescent intensity, (dIf/dt)o is the initial rate of gradual increase in intensity as solutes diffuse out of the vessel, and r is the venular radius. In each experiment, the venule was perfused under a constant perfusion pressure of 10 cmH2O and a flow velocity of 7 mm s-1. Vessels were discarded if fluorochrome leakage was detected.
Protein transfection of cells and venules
Protein transfection allows immediate studies of protein functions, avoiding the lag time of gene transcription and translation as required in conventional DNA transfection procedures. The transfection reagent TransIT-LT1 (Mirus) was used to promote the delivery of rVE-cad CPD to microvascular endothelial cells as well as intact microvessels. TransIT is composed of amphipathic polyamines and histone H1 protein which substantially enhance the transfection efficiency (
90%) of proteins and peptides, without affecting cell viability and function. The technical details and validation of the methodology have been provided in our previous publications (Tinsley et al. 1998, 2000, 2001; Yuan, 2000). Briefly, a cannulated venule was perfused at a constant perfusion pressure gradient of 20 cmH2O for 1 h with a transfection mixture containing the polyamine reagent TransIT-LT1 at 10 µl ml-1 and rVE-cad CPD at 1–5 µg ml-1. After transfection, the vessel was washed with a regular perfusate and then subjected to permeability measurements. The same procedure was used for monolayer transfection. As controls, cells or venules were transfected with the transfection reagent alone or TransIT-LT1 plus rVE-cad CPD mock extract; the latter was prepared by using the same procedure for rVE-cad CPD purification from E. coli cells without IPTG induction. For immunoprecipitation, immunoblotting and immunocytochemistry experiments, cells were harvested and processed at 2 h following transfection.
Data analyses
In the vessel studies, Pa was measured 2–3 times in each venule at each experimental intervention and the values were averaged. For all experiments n is given as the number of dishes of cells or vessels studied, with each vessel representing a separate animal. For each experiment, the Pa values obtained from different vessels or dishes of cells were averaged, normalized to the basal values obtained before treatment, and reported as percentage of basal value in mean ±S.E.M. ANOVA was used to evaluate the significance of intergroup differences. A P value of < 0.05 was considered significant for the comparison.
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Discussion |
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The organization and function of endothelial cadherin–catenin complexes in AJs are dynamically regulated by physical forces, cellular factors, and chemical mediators (Kusumi et al. 1999; Gumbiner, 2000; Bazzoni & Dejana, 2001; Dejana et al. 2001). During inflammation, an array of inflammatory agonists, including histamine, thrombin, growth factors, cytokines, and oxidants as well as activated leucocytes, can increase the transendothelial flux of fluid, macromolecules and cells across the microvascular wall through, to a large extent, the paracellular pathway (Leach et al. 1995; Bates & Curry, 1996, Bates & Harper, 2003; Del Maschio et al. 1996; Rabiet et al. 1996; Siflinger-Birnboim & Malik, 1996; Kevil et al. 1998; Andriopoulou et al. 1999; McDonald et al. 1999; Tinsley et al. 1999; Yuan, 2000, 2002; Dejana et al. 2001; Tiruppathi et al. 2001). Abnormalities in AJs accompanied by intercellular gap formation are often observed and provide a mechanistic basis for the paracellular leakage. Currently, two major hypotheses have been developed to explain the junctional changes in response to inflammation. One theory suggests that VE-cadherin and/or other junctional components may be degraded or subject to proteolytic cleavage when exposed to inflammatory agonists (Del Maschio et al. 1996; Allport et al. 1997; Bannerman et al. 1998; Carden et al. 1998; Moll et al. 1998; McDonald et al. 1999). This concept is challenged by the fact that the overall expression and protein mass of VE-cadherin are not reduced in endothelial cells during injury or when exposed to angiogenic stimulation (Gulino et al. 1998; Alexander et al. 2000; Wright et al. 2002), despite the occurrence of endothelial barrier dysfunction. The other hypothesis considers VE-cadherin endocytosis and sequestration as the key event in inflammation-induced AJ disorganization (Alexander et al. 2000; Gao et al. 2000). This is supported by experiments in which depleting extracellular calcium or blocking the extracellular domains of VE-cadherin with monoclonal antibodies leads to AJ disassembly and paracellular leakage (Corada et al. 1999, 2001; Alexander et al. 2000; Gao et al. 2000; Bazzoni & Dejana, 2001; Dejana et al. 2001). Apparently, disruption of VE-cadherin homophilic binding at the extracellular location serves as an effective means of altering the configuration of AJs. However, a question remains as to whether the changes in the extracellular configuration play an initial or key role in the regulation of vascular permeability especially under inflammatory conditions. In contrast, most inflammatory mediators act by binding to their respective receptors, which trigger cascades of intracellular signalling reactions that ultimately affect cellular functions. In this regard, the intracellular domain of the junctional protein possesses a higher potential than its extracellular counterpart to become the target of inflammatory signals.
The anchorage of VE-cadherin to the cytoskeleton is mediated by catenins, among which ß-catenin not only provides a structural linkage but also serves as a signalling molecule that coordinates the interaction between VE-cadherin and cytoskeletal elements (Yap et al. 1997; Gumbiner, 2000; Bazzoni & Dejana, 2001; Dejana et al. 2001; Venkiteswaran et al. 2002). We have previously reported that endothelial ß-catenin undergoes tyrosine phosphorylation in response to stimulation by C5a-activated neutrophils and that inhibition of the phosphorylation can greatly attenuate the increase in venular permeability caused by the same mediator (Tinsley et al. 1999, 2002). Interestingly, ß-catenin phosphorylation is accompanied by conformational changes at the cell–cell contacts characterized by diffuse VE-cadherin distribution and intercellular gap formation, the same morphology seen in the current study during disruption of VE-cadherin–catenin complexes. Notably, ß-catenin phosphorylation coupled with VE-cadherin disorganization has been observed in endothelial cells exposed to soluble permeability-enhancing agonists, including histamine, thrombin and vascular endothelial cell growth factor (Bates et al. 1996, 2003; Rabiet et al. 1996; Esser et al. 1998; Kevil et al. 1998; Andriopoulou et al. 1999; Cohen et al. 1999; Tiruppathi et al. 2001; Shasby et al. 2002). Therefore, it is possible that ß-catenin-signalled dissociation of VE-cadherin from its cytoskeletal anchor is involved in the transduction of the endothelial hyperpermeability response. In support of this hypothesis, the present study showed that competitive blockage of VE-cadherin-cytoskeleton binding resulted in a diffuse junctional organization concomitant with an increase in endothelial permeability. The data suggest that full-length VE-cadherin is required for the full strength of endothelial cell–cell adhesion. Furthermore, dissociation of VE-cadherin from its cytoskeletal anchor may play an important role in the mechanism responsible for inflammatory mediator-induced endothelial junctional disorganization and barrier dysfunction.
The functional importance of VE-cadherin intracellular domain is supported by previous studies showing that deletion or substitution of the cytoplasmic domains of cadherins blocks cell–cell aggregation (Nagafuchi & Takeichi, 1988; Ozawa et al. 1990; Kintner, 1992; Fujimori & Takeichi, 1993; Hermiston & Gordon, 1995; Hermiston et al. 1996; Zhu & Watt, 1996; Yap et al. 1997; Nieman et al. 1999). Targeted disruption or truncation of the ß-catenin binding sequence of VE-cadherin genes in mice induces endothelial apoptosis and impairs angiogenesis (Carmeliet et al. 1999; Gory-Faure et al. 1999). Cells transfected with VE-cadherin mutant lacking a C-terminal tail fail to form a tight barrier (Navarro et al. 1995). While these studies provide valuable information on the function of the cadherin cytoplasmic domain, most of them were conducted in cells that do not participate in the microvascular exchange process. Furthermore, DNA transfection and gene manipulation are often subject to technical problems associated with low transfection efficiency, inconsistency, and transcriptional or translational variations (Kotnis et al. 1995; Teifel et al. 1997). It is worth noting that our study used a different approach. We expressed and purified a recombinant fusion protein of VE-cadherin cytoplasmic domain as a molecular tool to specifically block the binding of endogenous VE-cadherin to ß-catenin. The protein was delivered into the intact microvascular endothelium using a novel protein transfection technique recently developed in our laboratory (Tinsley et al. 1998, 2000, 2001; Yuan, 2000). The polyamine-based transfection method efficiently delivers peptides and proteins into the vascular endothelial cells as well as the intact microvascular wall without any significantly compromising cell viability or vessel functions (Tinsley et al. 1998, 2000, 2001; Yuan, 2000). In this study, rVE-cad CPD was observed inside the cytoplasm of transfected cells by using immunofluorescence microscopy, indicating the efficient delivery of this protein into cells. The recombinant protein interacted with ß-catenin to form complexes in vitro and in vivo and competed with endogenous VE-cadherin for ß-catenin binding in cells, as indicated by an attenuated binding interaction between the two molecules. The competition was further evidenced by the concomitant increase in rVE-cad CPD-ß-catenin binding and decrease in VE-cadherin–ß-catenin binding. This was also supported by the data showing a decreased partitioning of VE-cadherin in the detergent-insoluble pool. Note that we do not exclude the possibility that the binding of cytoplasmic ß-catenin to rVE-cad CPD could modify the amount of nuclear ß-catenin as a consequence of its transcriptional activity, inducing a more complex effect that indirectly affects the junctional composition. However, it is not likely that a relatively long-term effect of transcriptional modulation was responsible for the acute change in the venular permeability. The early occurrence of hyperpermeability in the intact venules following rVE-cad CPD perfusion supports the direct effect of rVE-cad CPD on VE-cadherin composition. To the best of our knowledge, this is the first report on specific interruption of the binding between VE-cadherin and ß-catenin and its impact on microvascular permeability.
In conclusion, this study reveals the functional impact of disrupting the structural association between VE-cadherin and ß-catenin-linked cytoskeleton on endothelial junctional integrity. Transfection of rVE-cad CPD into the microvascular endothelium, which competes with endogenous VE-cadherin for binding to ß-catenin, results in junctional disorganization and venular hyperpermeability. We suggest that the intracellular association of VE-cadherin with cytoskeletal proteins is of critical importance in the regulation of microvascular barrier function.
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catenins, respectively. B, schematic composition of the expression vector construct pQE30/VE-cad-CPD. PT5, phage promoter T5; lacO, lac operator; RBS, ribosome binding site; ATG, start codon; 6xHis, histidine tag. C, cDNA product of VE-cadherin cytoplasmic domain on agarose gel stained with ethidium bromide. D, Western blot detection of the rVE-cad CPD protein. Lane 1, purified protein; lane 2, mock extract from E. coli without IPTG induction; lanes 3–5, cell lysate from different bacterial clones.
