Identification of Proteases Involved in the Proteolysis of Vascular Endothelium Cadherin during Neutrophil Transmigration*

Bastien HermantDagger §, Stéphanie BibertDagger , Evelyne ConcordDagger , Bernard Dublet, Marianne WeidenhauptDagger ||, Thierry VernetDagger , and Danielle Gulino-DebracDagger **

From the Dagger  Laboratoire d'Ingénierie des Macromolécules and  Laboratoire de Spectrométrie de Masse des Protéines, Institut de Biologie Structurale Jean-Pierre Ebel (Commissariat à l'Energie Atomique/CNRS, Université Joseph Fourier), 41 rue Jules Horowitz, 38027 Grenoble Cedex, France

Received for publication, January 13, 2003, and in revised form, February 7, 2003

    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Transmigration of neutrophils across the endothelium occurs at the cell-cell junctions where the vascular endothelium cadherin (VE cadherin) is expressed. This adhesive receptor was previously demonstrated to be involved in the maintenance of endothelium integrity. We propose that neutrophil transmigration across the vascular endothelium goes in parallel with cleavage of VE cadherin by elastase and cathepsin G present on the surface of neutrophils. This hypothesis is supported by the following lines of evidence. 1) Proteolytic fragments of VE cadherin are released into the culture medium upon adhesion of neutrophils to endothelial cell monolayers; 2) conditioned culture medium, obtained after neutrophil adhesion to endothelial monolayers, cleaves the recombinantly expressed VE cadherin extracellular domain; 3) these cleavages are inhibited by inhibitors of elastase; 4) VE cadherin fragments produced by conditioned culture medium or by exogenously added elastase are identical as shown by N-terminal sequencing and mass spectrometry analysis; 5) both elastase- and cathepsin G-specific VE cadherin cleavage patterns are produced upon incubation with tumor necrosis factor alpha -stimulated and fixed neutrophils; 6) transendothelial permeability increases in vitro upon addition of either elastase or cathepsin G; and 7) neutrophil transmigration is reduced in vitro in the presence of elastase and cathepsin G inhibitors. Our results suggest that cleavage of VE cadherin by neutrophil surface-bound proteases induces formation of gaps through which neutrophils transmigrate.

    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The endothelium constitutes a physical barrier between blood and the underlying tissues and is the point of entry of leukocytes into inflamed tissues. The mechanism of leukocyte entrance into tissues is a complex, multistep event involving leukocyte adhesion to endothelium and subsequent migration across the blood vessel wall (1). Whereas the early steps leading to leukocyte adhesion are well understood (2, 3), it is still unclear how leukocytes crawl through the endothelial wall. Many studies demonstrated that leukocytes exit blood vessels by squeezing between adjacent endothelial cells (4), but a recent study suggested that leukocytes could pass across the body of endothelial cells using a transcellular route (5).

Two adhesive receptors, platelet-endothelial cell adhesion molecule-1 (PECAM-1)1 (6, 8, 9) and CD99 (7), were clearly demonstrated to be involved in the leukocyte transmigration process. They are both expressed at the surface of leukocytes and at endothelial cell-cell junctions and elaborate homophilic interactions between the two cellular types. In fact, PECAM-1-mediated homotypic interactions are relayed by CD99-mediated ones to regulate the transmigration of leukocytes across the endothelium.

Additionally, a third adhesive receptor, VE cadherin (10), expressed at endothelial cell-cell junctions, can modulate leukocyte transmigration. VE cadherin participates in the elaboration of inter-endothelial adherens junctions as its extracellular part establishes homophilic interactions resulting in cell-cell attachment (11, 12). These ectodomain-based interactions are reinforced by interactions involving the cytoplasmic domain of VE cadherin, which binds, in a mutually exclusive manner, the armadillo proteins beta  or gamma  catenin (13, 14). By mediating alpha  catenin binding (15), F-actin-bundling proteins, beta  or gamma  catenins, promote connections between the actin cytoskeleton and the VE cadherin-based intercellular junction (16). An other member of the armadillo family, p120ctn or delta  catenin, binds to the membrane proximal region of the VE cadherin cytoplasmic part and modulates the adhesive strength of VE cadherin (17). By establishing these multiple interactions, VE cadherin plays an important role in the maintenance of the endothelial integrity (18). Indeed, anti-VE cadherin antibodies, when administered to mice, induce an increase in vascular permeability (19) and, when added to endothelial cell monolayers, provoke the appearance of gaps at cell-cell junctions (18).

VE cadherin also possesses a critical role in the transmigration of leukocytes. Indeed, treatment with anti-VE cadherin antibodies, able to disrupt interendothelial junctions, results in an increase of neutrophil transmigration in vitro (20) and in vivo (19, 44). Moreover, immunofluorescence staining reveals that VE cadherin and alpha , beta , and gamma  catenins disappear from cell-to-cell contacts following neutrophil adhesion to endothelial monolayers. This effect appears only where neutrophils firmly adhere, whereas VE cadherin-based complexes remain intact in areas devoid of adherent neutrophils (21, 22). By contrast, PECAM-1 distribution remained unaffected (22).

The exact role played by VE cadherin in the transmigration process is still controversial. It was first proposed that leukocyte adhesion might trigger intracellular signals, leading to the destabilization of VE cadherin-based complexes and thus allowing leukocyte transmigration to occur through the formed gaps (22, 21). Indeed, disruption of the VE cadherin/catenin complexes (23) generally parallels with the tyrosine phosphorylation of the different partners within the cadherin complex. These phosphorylation events may be modulated by the capacity of VE cadherin to associate with kinases such as the receptor tyrosine kinase VEGFR-2 (24, 25) and with phosphatases such as vascular endothelial protein tyrosine phosphatase (VE-PTP) (26) and SH 2 domain-containing tyrosine phosphatase 2 (SHP-2) (27, 28).

A second and more recent hypothesis concerning the role of VE cadherin in transmigration proposes that migrating leukocytes locally push VE cadherin in the plane of the junction thus allowing their passage through the intercellular gaps (4). In this study, transmigration of fluorescently labeled leukocytes was followed in real time by two-color fluorescence microscopy using HUVECs expressing endogenous VE cadherin and a VE cadherin protein fused in its C-terminal part with green fluorescent protein. This study shows that transmigration occurs through both preexisting gaps and through de novo gap formation. Gap widening accommodates to the size of the transmigrating leukocyte allowing a narrow contact between endothelial cell and leukocyte. This widening of the gaps seems to be accompanied by a clustering of the VE cadherin protein fused in its C-terminal part with green fluorescent protein molecules on the edge of the forming clefts. After transmigration, the displaced molecules diffuse back to reconstitute the junctions. These results were recently confirmed by a real-time study following the differential movements of VE cadherin in living endothelial cells during transmigration of neutrophils using antibodies which do not interfere with transendothelial migration (29). The lateral movement of VE cadherin was hypothesized to be a consequence of its decoupling from the cytoskeleton, probably initiated by an intracellular signal as a result of leukocyte adhesion to the endothelium.

A possible role for neutrophil proteases is also suspected in transmigration (30). These proteases could cleave proteins participating in the elaboration of intercellular contacts, thus facilitating the progression of neutrophils across the endothelial junctions. Among the proteases susceptible to involvement in transmigration are serine proteases (including elastase, cathepsin G, and proteinase 3) and matrix metalloproteases (such as MMP-8 and MMP-9), all packed in the azurophil granules of neutrophils (31). Recently, elastase was suspected to be able to degrade components of VE cadherin complexes during contact between neutrophils and endothelial cells (22, 21), but this result remains controversial. Indeed, the adhesion-dependent degradation of endothelial catenins described by Del Maschio et al. (22) and Allport et al. (21) probably does not reflect transmigration events but rather a release of neutrophil proteases upon cell lysis (32). Moreover, the fact that, in MMP-9- and neutrophil elastase-deficient mice, neutrophils transmigrate as efficiently as in wild-type mice suggests that elastase and MMP-9 are not essential for the leukocyte transmigration process (33). This is in conflict with the results of Cepinskas et al. (30), who found that, on migrating neutrophils, membrane-bound elastase, which localizes to the migrating front, facilitates transendothelial migration. Moreover, in some pathological situations, administration of an elastase inhibitor attenuates the transmigration of neutrophils (34).

In the present report, we study the role of proteases and VE cadherin in neutrophil transmigration. We demonstrate that, following adhesion of neutrophils to endothelial monolayers, VE cadherin fragments are released from the surface of endothelial cells, suggesting that proteases are involved in the transmigration process. We present evidence that purified elastase and cathepsin G can cleave in vitro the extracellular part of VE cadherin. The exact positioning of the cleavage sites for each protease allows us to clearly identify elastase and cathepsin G as the surface-bound proteases able to cleave VE cadherin. Moreover, we also demonstrate that specific blockage of both elastase and cathepsin G at the neutrophil surface significantly reduces their transmigration in an in vitro assay. This strongly suggests that proteolytic events such as the cleavage of VE cadherin occurring at cell-cell junctions are required for neutrophil transmigration.

    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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Cells-- Neutrophils were isolated from the peripheral blood of healthy volunteers by using dextran-Ficoll (35) and resuspended in M199 medium (Biowhittaker, Verviers, Belgium). Human endothelial cells were isolated from umbilical veins and cultured in M199 supplemented with 20% fetal bovine serum (Biowhittaker), 50 µg/ml endothelial cell growth supplement (Biowhittaker) and kept at 37 °C in a 5% CO2 humidified atmosphere, as previously described (10).

Reagents-- Elastase was purchased from Athens Research (Athens, GA); cathepsin G was from Elastin Products Co., Inc. (Owensville, MI); human Secretory leukocyte protease inhibitor (SLPI) was from R&D Systems Europe (Abingdon, UK); the specific human leukocyte elastase inhibitor N-methoxysuccinyl-Ala-Ala-Pro-Val chloromethyl ketone (MeOSuc-AAPV) was from Sigma-Aldrich Chimie (St Quentin Fallavier, France). Human tumor necrosis factor alpha  (TNFalpha ) and N-formyl-Met-Leu-Phe (fMLP) were from Roche Molecular Biochemicals (Meylan, France) and Sigma, respectively.

The bacterially expressed VE-EC1-4 fragment was produced as previously described (11, 12).

Antibodies-- Four fragments, designated as EC1, EC4, Cad1, and Cad3, encompassing various regions of the VE cadherin protein were used as antigens to raise the anti-VE cadherin polyclonal antibodies named anti-EC1, anti-EC4, anti-Cad1, and anti-Cad3, respectively. The shorter fragments VE-EC1 and VE-EC4 were produced as described (11). The cDNA constructs encoding Cad1 and Cad3 were elaborated as described previously for VE cadherin (18). The resulting plasmids expressed the VE cadherin fragments Cad1 and Cad3 (amino acid stretches 1-439 for Cad1 and 259-439 for Cad3) fused at their N termini with a 16-amino acid fusion peptide (18).

Production of the polyclonal anti-VE cadherin antisera was performed as previously described (18). Each polyclonal antibody was directly purified by applying the corresponding antiserum on an affinity column coupled to the recombinant fragment used as an antigen. The characterization of these polyclonal antibodies was performed by fluorescence-activated cell sorter analysis on Chinese hamster ovary cells expressing human VE cadherin, by immunofluorescence staining and Western blotting as previously described (18).

The anti-fragment antibodies were biotinylated with D-biotinoyl-epsilon -aminocaproic acid N-hydroxysuccimide ester according to the instructions from the supplier (Roche Molecular Biochemicals).

Capture Immunoassays-- Polyvinyl microtiter wells were coated overnight at 4 °C with 100 µl of either anti-EC1 or anti-EC4 or anti-Cad3 antibodies diluted at 10 µg/ml in 0.1 M sodium carbonate buffer, pH 9.6. The plates were postcoated with phosphate-buffered saline containing 1% bovine serum albumin, washed with 0.05 M phosphate-buffered saline plus 0.05% Tween 20, and incubated with 100 µl of cell supernatants overnight at 4 °C. Following incubation with supernatants, extensive washing was performed with 0.05 M phosphate-buffered saline, 0.05% Tween 20. Bound VE cadherin-derived fragments were detected by adding 100 µl of either biotinylated anti-Cad 1 or biotinylated anti-Cad 3 diluted in phosphate-buffered saline containing 0.5% bovine serum albumin and incubation was performed for 2 h at room temperature. After washing with phosphate-buffered saline and 0.05% Tween 20, alkaline phosphatase-conjugated streptavidin was added for 1 h at room temperature. After washing, binding was quantified using p-nitrophenyl phosphate as a substrate for alkaline phosphatase and measuring the absorbance at 405 nm.

Western Blotting-- Following its proteolysis, VE-EC1-4 was subjected to 12.5% polyacrylamide gel electrophoresis. Separated fragments were then electroblotted onto pure nitrocellulose membranes (Bio-Rad, Marnes-la-Coquette, France) and blocked with 3% gelatin-containing PBS. The blots were incubated overnight with the polyclonal anti-Cad3 antibody in PBS containing 1% gelatin, and protein bands were then detected by a peroxidase-labeled goat anti-rabbit antibody using the ECL kit (Amersham Biosciences).

Neutrophil Adhesion to Endothelial Cells-- Prior to the addition of neutrophils, confluent endothelial cell monolayers were treated with 100 units/ml TNFalpha for 24 h and then washed with PBS. Neutrophils, suspended in M199 medium containing 10 µM fMLP, were then added to monolayers at final leukocyte:endothelial cell ratios of either 10:1 (Fig. 3) or 4:1 (Figs. 4 and 5) and incubated for 1 h at 37 °C. Under this condition, the integrity of endothelial cell monolayers was preserved. In some experiments, neutrophils were layered on resting endothelial cells. Following an incubation of 1 h, cell culture media containing nonadherent neutrophils were collected and gently centrifuged (300 × g, 10 min) to avoid neutrophil crushing. Supernatants were separated from neutrophil cells and then tested for their capacity to cleave the recombinant fragment VE-EC1-4.

Digestion of the Recombinant Fragment VE-EC1-4 by Elastase, Cathepsin G, or Cell Medium Supernatant-- VE-EC1-4 (final concentration, 3 µM) was mixed with either elastase (final concentration, 0.02 unit/ml) or cathepsin G (final concentration, 0.02 unit/ml) or the supernatant (dilution 1/3, see above). The mixtures were incubated for different periods of time at 37 °C prior to Western blot or matrix-assisted laser desorption (MALDI) mass spectrometry analyses. In some assays, the purified proteases or the supernatants were preincubated with the protease inhibitors MeOSuc-AAPV or with SLPI at 37 °C during 20 min before addition of the fragment VE-EC1-4.

Degradation of the Recombinant Fragment VE-EC1-4 by Cell Surface-bound Enzymes-- Neutrophils were stimulated for 30 min at 37 °C both with fMLP (10-8 M) and TNFalpha (100 units/ml) as previously described (36). Following stimulation, neutrophils were fixed for 3 min at 4 °C with PBS containing 3% paraformaldehyde (w/v) and 1% glutaraldehyde (v/v). Neutrophils were then centrifuged at 300 × g for 5 min, washed three times in PBS, and resuspended in cell culture medium M199 without serum. Either fixed neutrophils (105 cells) or purified human leukocyte elastase (7 × 10-4 units) or purified human leukocyte cathepsin G (7 × 10-4 units) were added to the recombinant fragment VE-EC1-4 (5 µg) in the presence or absence of either SLPI (10-10 mol) or MeOSuc-AAPV (10-10 mol) and incubated at 37 °C for 3 h. Following centrifugation at 300 × g for 5 min, the cell-free supernatants were mixed with beta -mercaptoethanol containing Laemmli buffer and subjected to SDS-polyacrylamide electrophoresis. Western blots were then performed as described above.

Transendothelial Permeability Assay-- Cells were cultured until confluence on polycarbonate membranes of Transwell inserts (0.4-µm pore size, Costar, Cambridge, MA) as previously described (18). Following verification of cell confluence by crystal blue staining, the culture medium in the upper compartment was substituted with medium containing purified elastase (2 × 10-2 units/ml), purified cathepsin G (2 × 10-2 units/ml), or cell culture supernatant (see above). At the same time, horseradish peroxidase-linked goat IgG (Rockland, Gilberville, PA) was added to the upper compartment of each Transwell unit. After various incubation times, the medium in the lower compartments was assayed photometrically for the presence of peroxidase with o-phenyldiamine dihydrochloride (Dako, Glostrup, Denmark) as a substrate according to the instructions from the supplier. Three individual Transwell units were used for each incubation time. To verify whether protease inhibitors modify endothelial permeability, SLPI (4 × 10-10 mol/Transwell insert) or MeOSuc-AAPV (2 × 10-6 mol/Transwell insert) were added to the upper compartments of Transwell units and the kinetic of marker migration across endothelial monolayers was established as described above.

MALDI Mass Spectrometry-- Mass spectra of the proteolytic fragments derived from VE-EC1-4 were obtained with a Perspective Biosystems Voyager Elite Xl time of flight mass spectrometer with delayed extraction, operating with a pulsed nitrogen laser at 337 nm (Framingham, MA). Positive-ion mass spectra were acquired using linear, delayed extraction mode with an accelerating potential of 25 kV, a 90% grid potential, a 0.2% guide wire voltage, and a delay time of 200 ns. Each spectrum is the result of 200 averaged laser pulses.

Samples were mixed with an equal volume of 1% trifluoroacetic acid and concentrated on a ZipTipTM C4 (Millipore) as specified by the manufacturer. The material eluted from the ZipTipTM C4 was partially evaporated and mixed with an equal volume of a saturated solution of sinapinic acid prepared in 50% (v/v) aqueous acetonitrile, 0.1% trifluoroacetic acid. Aliquots of 2 µl of this mixture were spotted on the stainless steel sample plate and dried in the air prior to analysis. External calibration was performed with enolase from bakers' yeast using the m/z values of 46,672 and 23,336 for the mono- and di-charged molecules, respectively. The accuracy of MALDI molecular weight determinations comprises between 0.01 and 0.2%.

Blotting for N-terminal Sequencing-- Following electrophoresis, proteolytic products were transferred to polyvinylidene difluoride ProblottTM membranes (Applied Biosystems, Foster City, CA) using a 10 mM CAPS, 10% (v/v) methanol buffer at pH 11. Transfer was carried out during 1 h at room temperature using a constant voltage of 50 V. After transfer, the membranes were rinsed with distilled water, saturated with 100% methanol for a few seconds, stained for 5 min with a solution of 0.1% Coomassie Blue R250 in acetic acid/methanol/H2O (10:40:50, v/v/v), and then destained with 50% methanol and air-dried. Protein bands were excised and submitted to N-terminal sequencing. Amino acid sequence determination was performed using a gas-phase Sequencer model 477A (Applied Biosystems).

Neutrophil Transmigration Assay-- HUVECs were grown to confluence on fibronectin-coated porous membranes of Transwell units (3-µm pore size, Falcon, Dutscher, Issy-les-Moulineaux, France) for 3 days at 37 °C. To study the influence of proteases on transmigration, freshly purified neutrophils were activated at 37 °C during 30 min in M199 containing 20% fetal bovine serum, 100 units/ml TNFalpha , and 10-8 M fMLP. In these conditions, proteases are translocated from azurophil granules to the external surface of neutrophils (36). In some assays, to inhibit surface proteases, activated neutrophils were incubated with either SLPI (2.5 × 10-10 mol/assay) or MeOSuc-AAPV (2 × 10-6 mol/assay) during 2 h at 37 °C. Then, activated neutrophils, treated or not with inhibitors, were rinsed, washed twice in PBS, and diluted in M199 medium containing 20% fetal bovine serum before their addition to TNFalpha -activated (5 × 104 cells/insert). To create a chemotactic gradient for neutrophils, 2 × 10-8 M fMLP in M199 was added to the lower compartments of the Transwell units. To evaluate the number of transmigrated neutrophils, the Transwell units devoid of their upper compartments were centrifuged at 4 °C for 10 min at 300 × g. The supernatants were discarded and the cell pellet lysed in experimental conditions allowing the extraction of neutrophil DNA using the high pure template kit (Roche Molecular Biochemicals). The number of transmigrated neutrophils was expressed as a function of the amount of neutrophil DNA, quantified by measuring optical density at 260 nm.

In parallel, the presence of active proteases at the surface of activated neutrophils or their inactivation following treatment with inhibitors was verified by incubating differently treated neutrophils with the fragment VE-EC1-4 as described above. As indicated by blue trypan staining, the various treatments applied to neutrophils did not induce cell death.

    RESULTS
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INTRODUCTION
MATERIALS AND METHODS
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Cleavage of VE Cadherin Induced by Adhesion of Neutrophils onto Endothelial Cell Monolayers-- To investigate the fate of VE cadherin during transmigration of leukocytes across endothelium, human neutrophils were added, in the presence of the chemoattractant reagent fMLP, to confluent TNFalpha -activated endothelial cell monolayers. In these conditions, neutrophils could adhere onto endothelial cells. Following this adhesion, we observe VE cadherin fragments in the cell culture supernatants using an immunocapture assay (Fig. 1). As indicated in Fig. 1A, the amount of soluble VE cadherin fragments increased as contact time between endothelial cells and neutrophils increased. Moreover, the level of VE cadherin accumulated in the cell culture supernatant could be correlated with the neutrophil/HUVEC ratio (Fig. 1B).


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Fig. 1.   Detection of soluble VE cadherin-derived fragments in cell culture supernatants following adhesion of neutrophils on endothelial cell monolayers. A, increase of the amount of VE cadherin fragments with contact time between endothelial cells and neutrophils. Neutrophils were incubated on TNFalpha -activated HUVEC monolayers during 0, 1, or 2 h at 37 °C using a neutrophil/HUVEC ratio of 10. The immunocapture assay was performed using the antibodies anti-EC1 for coating and biotinylated anti-Cad3 for detection of soluble VE cadherin. B, increase of the amount of soluble VE cadherin with increasing neutrophil/HUVEC ratio. Neutrophils were incubated on TNFalpha -activated HUVEC monolayers during 1 h at 37 °C using neutrophil/HUVEC ratios of 0, 10, and 15. Soluble VE cadherin fragments were detected using the immunocapture assay described in panel A. In panels A and B, bars at the left correspond to the background absorption.

To understand the mechanism underlying the production of soluble VE cadherin fragments, an ELISA using the antibodies anti-EC1 and anti-Cad3 (Fig. 2A) was performed on cell culture supernatants obtained from differentially treated endothelial cells (Fig. 2B, ELISA 1). First, to analyze the role of TNFalpha in this process, the content of soluble VE cadherin in supernatants derived from endothelial cells either activated or not by this cytokine was quantified. Slightly more soluble VE cadherin fragments were detected in TNFalpha -activated supernatants when compared with supernatants from untreated cells. To test the impact of neutrophil adhesion to endothelial cells, the content of VE cadherin fragments in supernatants from TNFalpha -activated endothelial cells on which neutrophils had adhered was measured. Results showed that adhesion of neutrophils to TNFalpha -activated cells significantly increased the amount of VE cadherin detected in the supernatants. In contrast, a very low amount of VE cadherin fragments was detected in supernatants derived from cells on which neutrophils had not adhered (i.e. in assays in which neutrophils were layered on resting endothelial cells).


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Fig. 2.   Relationship between adhesion of neutrophils on endothelial cell monolayers and accumulation of soluble fragments of VE cadherin in cell culture supernatants. A, schematic representation of VE cadherin and recombinant fragments used to raise the polyclonal antibodies anti-Cad1, anti-EC1, anti-EC4, and anti-Cad3. The black bar in Cad1 and Cad3 represents an N-terminal fusion peptide. B, detection of soluble fragments of VE cadherin using various ELISAs. Neutrophils were added to either TNFalpha -activated (black) or non-activated (light gray) endothelial cell monolayers. After a 1-h contact period at 37 °C, supernatants were collected and their content in VE cadherin-derived fragments analyzed using ELISA. For comparison, supernatants derived from TNFalpha -activated (dark gray) and non-activated (white) endothelial cell monolayers without neutrophil treatment were also collected and analyzed in parallel. ELISAs were performed using five different sets of polyclonal antibody pairs; capture of VE cadherin fragments was obtained with different polyclonal anti-VE cadherin antibodies (1 and 2, anti-EC1; 3 and 4, anti-Cad3; 5, anti-EC4); detection of the bound fragments was then performed using various biotinylated anti-VE cadherin antibodies (1, 3, and 5, anti-Cad3; 2 and 4, anti-Cad1).

In the ELISA 1, depicted in Fig. 2B, only fragments overlapping at least the amino acid stretch Phe-104 to Phe-259 and extending maximally from amino acid Asp-1 to Lys-439 of the VE cadherin sequence could be detected. To improve the detection of VE cadherin-derived fragments, several ELISAs were performed using different pairs of polyclonal antibodies directed against various regions of VE cadherin (Fig. 2A). In each case, adhesion of neutrophils onto endothelial cells resulted in a significant increase of the level of soluble fragments into cell supernatants (Fig. 2B, ELISA 2-5). This suggested that adhesion of neutrophils to TNFalpha -activated endothelial cells induces the cleavage of the extracellular part of VE cadherin.

In Vitro Cleavage of the VE Cadherin Extracellular Region by Cell Culture Supernatants-- We hypothesized that adhesion of neutrophils to TNFalpha -activated endothelial monolayers activated neutrophil proteases, which could release extracellular fragments of VE cadherin from the cell surface into the culture medium.

In an attempt to detect the proteases able to cleave VE cadherin, the cell culture supernatants were mixed with a recombinant protein encompassing the majority of the extracellular part of VE cadherin, designated as VE-EC1-4, which was previously demonstrated to be able to self-assemble as a stable hexamer (11, 12).

Treatment of endothelial cells with either TNFalpha or fMLP or both gave supernatants that did not induce a significant degradation of the recombinant fragment (Fig. 3, lanes 2-4). Moreover, supernatants obtained after addition of neutrophils to non-activated endothelial cells left the recombinant fragment intact, indicating that no active protease was present in this condition (Fig. 3, lane 5). In contrast, a complete degradation of the fragment was observed with supernatants derived from endothelial cells on which neutrophils had adhered (Fig. 3, lane 6). This suggested that adhesion of neutrophils on endothelial cells caused a secretion and/or an activation of proteases in the cell culture supernatant.


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Fig. 3.   Supernatant-induced proteolysis of the recombinant fragment VE-EC1-4. The recombinant fragment VE-EC1-4 (10 µM) was mixed with endothelial cell supernatants and incubated at 37 °C for 1 h. The mixtures were electrophoresed on a 7.5% polyacrylamide gel and transferred on nitrocellulose membranes before probing with the polyclonal antibody anti-Cad3 (Fig. 2A). Supernatants were obtained from TNFalpha -treated (lanes 2, 4, and 6) or non-treated (lanes 1, 3, and 5) endothelial cells in the presence (lanes 3, 4, and 6) or in the absence (lanes 1, 2, and 5) of fMLP. Lanes 5 and 6 correspond to supernatants derived from endothelial cells to which neutrophils had been added (neutrophils/endothelial cell ratio = 10).

To observe the potential intermediate proteolytic fragments, the neutrophil:endothelial cell ratio was diminished from 10:1 to 4:1 and cell culture supernatants were diluted (1/3) prior to their addition to the recombinant fragment VE-EC1-4. The time course of the proteolysis was followed by Western blot using the polyclonal anti-VE cadherin anti-Cad3 antibody directed against the C-terminal part of VE-EC1-4. In these conditions, two distinct cleavage products with apparent molecular masses of 38 and 27 kDa were detected 2 or 4 h after addition of the culture supernatant to VE-EC1-4 (Fig. 4A). Re-probing the Western blot membranes with the anti-VE cadherin antibody directed against the N-terminal module EC1 (Fig. 2A) allowed the detection of an additional fragment of 22 kDa (data not shown). These results were systematically observed using neutrophils from 10 different donors (data not shown).


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Fig. 4.   Nature of the supernatant-containing proteases. Supernatants used in panels A and B of this figure were obtained in conditions identical to those depicted in Fig. 3 (lane 6). The molecular masses of the markers are indicated at the left margin in kDa. A, time course of supernatant-induced proteolysis of VE-EC1-4. Supernatants were diluted to 1/3 and mixed with VE-EC1-4 (final concentration, 3 µM) (lanes 2, 4, and 6). In control experiments, buffer was mixed with VE-EC1-4 (lanes 1, 3, and 5). The mixtures were then incubated at 37 °C during different periods of time (0 h, lanes 1 and 2; 2 h, lanes 3 and 4; 4 h, lanes 5 and 6). B, supernatant-induced proteolysis is blocked by the protease inhibitor SLPI. VE-EC1-4 (3 µM) was mixed either with supernatant (dilution 1/3) (lanes 4-9) or, in controls, with buffer (lanes 1-3). The various mixtures were incubated at 37 °C during 0 h (lanes 1, 4, and 7), 2 h (lanes 2, 5, and 8), and 4 h (lanes 3, 6, and 9). SLPI was added at 0.5 µg/ml to the mixtures (lanes 7, 8, and 9). C, VE-EC1-4 digestion by purified elastase. VE-EC1-4 (3 µM) was mixed either with purified elastase (0.02 unit/ml) (lanes 4-9) or, in controls, with buffer (lanes 1-3). The various mixtures were incubated at 37 °C during 0 h (lanes 1, 4, and 7), 2 h (lanes 2, 5, and 8), and 4 h (lanes 3, 6, and 9). SLPI was added at 0.5 µg/ml to the mixtures (lanes 7-9). D, VE-EC1-4 digestion by purified cathepsin G, VE-EC1-4 (3 µM) was mixed either with purified cathepsin G (0.02 unit/ml) (lanes 4-9) or, in controls, with buffer (lanes 1-3). The various mixtures were incubated at 37 °C during 0 h (lanes 1, 4, and 7), 2 h (lanes 2, 5, and 8), and 4 h (lanes 3, 6, and 9). SLPI was added at 0.5 µg/ml to the mixtures (lanes 7-9). Western blot analysis was then performed as described in Fig. 3.

Identification of the Neutrophil Proteases Involved in the Cleavage of VE Cadherin-- To identify the supernatant-containing proteases involved in the VE cadherin cleavage, we used a physiological inhibitor, the human SLPI, in our cleavage assays. SLPI is known to specifically block the neutrophil granule-containing proteases elastase and cathepsin G (37). At a final concentration of 0.5 µg/ml, SLPI effectively blocked the supernatant-induced degradation of VE-EC1-4 (Fig. 4B, lanes 7-9). This suggested strongly that elastase and/or cathepsin G were the active proteases present in the supernatants.

We then verified that purified elastase, at concentrations as low as 0.02 unit/ml, was able to significantly cleave VE-EC1-4. The pattern of digestion of this enzyme (Fig. 4C, lanes 5 and 6) appeared very similar to those obtained with the supernatants (Fig. 4B, lanes 5 and 6). Indeed, both the 38- and the 27-kDa proteolytic products were clearly detected by Western blot following 2 or 4 h of treatment. This strongly indicated that one of the proteases secreted by neutrophils, following their adhesion on endothelial cells, is elastase. Similarly, purified cathepsin G was also able to cleave VE-EC1-4. After a 2-h incubation time, the digestion pattern (Fig. 4D, lane 5) was very similar to that observed with purified elastase (Fig. 4C, lane 5). In contrast, 4 h after addition of cathepsin G, the 38-kDa band disappeared, whereas the 27-kDa band appeared slightly less abundant (Fig. 4D, lane 6). Furthermore, addition of SLPI to each purified enzyme blocked their proteolytic activity. Indeed, as shown in Fig. 4 (C and D, lanes 8 and 9), SLPI inhibited the elastase activity totally and that of cathepsin G partially.

To confirm that elastase was one of the proteases responsible for the proteolytic cleavage of fragment VE-EC1-4, the chemical neutrophil elastase-specific inhibitor MeOSuc-AAPV (37) was incubated with supernatant before its addition to VE-EC1-4. In these conditions, intensities of both the 38- and 27-kDa proteolytic bands were strongly decreased (Fig. 5, lanes 7-10). Concomitantly, we also verified that this elastase-specific inhibitor only affects the elastase activity (Fig. 5, lanes 3-6) and not the cathepsin G activity (Fig. 5, lanes 11-14).


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Fig. 5.   Presence of elastase within cell supernatants. Supernatants used in this figure were obtained in conditions identical to those depicted in Fig. 3 (lane 6). VE-EC1-4 (3 µM) was mixed either with purified elastase (0.02 unit/ml) (lanes 3-6) or with purified cathepsin G (0.02 unit/ml) (lanes 11-14) or with supernatant (dilution 1/3) (lanes 7-10) or, in control assays, with buffer (lanes 1 and 2) and incubated at 37 °C during 0 h (lanes 1, 3, 5, 7, 9, 11, and 13) or 2 h (lanes 2, 4, 6, 8, 10, 12, and 14). The specific inhibitor of elastase MeOSuc-AAPV was added at a final concentration of 0.5 mM (lanes 5, 6, 9, 10, 13, and 14). Detection of the proteolytic fragments was performed as described in Fig. 4.

Identification of the Elastase- and Cathepsin G-induced Cleavage Sites by N-terminal Sequencing and MALDI Mass Spectrometry-- N-terminal sequencing of the 27-kDa electroblotted bands observed in Fig. 4 (panels C and D, lanes 5) showed that they corresponded to C-terminal peptides beginning at positions Thr-201 or Gln-203 when VE-EC1-4 was digested either by elastase or cathepsin G, respectively (Table I). The 38-kDa band generated by elastase digestion corresponded to a single C-terminal peptide starting at the amino acid Lys-94. In contrast, the 38-kDa cathepsin G-generated band contained two peptides starting at amino acids Thr-92 and Val-95 (Table I).


                              
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Table I
Identification by N-terminal sequencing and MALDI mass spectrometry of the proteolytic peptides derived from VE-EC1-4 digested with either purified elastase or purified cathepsin G or cell supernatant
Fragment VE-EC1-4 was mixed with either elastase or cathepsin G or cell supernatant as previously described in Fig. 6. Following proteolysis, the various mixtures were first analyzed by MALDI mass spectrometry. In parallel, N-terminal sequencing was performed on the proteolytic fragments previously electroblotted onto polyvinylidene difluoride membrane. down-arrow indicates the position of the cleavage site. The calculated molecular weights of the proteolytic fragments were determined using the computer program Protparam Tools of the Expasy server (www.expasy.ch/tools/protparam.html). N.D., not determined.

MALDI mass spectrometry analyses were also performed to confirm and complete the N-terminal sequencing data (Fig. 6). Mass spectra indicated that treatment of VE-EC1-4 with purified elastase gave a major proteolytic product possessing a molecular mass of 26,611 Da (Fig. 6A and Table I), whereas purified cathepsin G released a product of 26,424 Da (Fig. 6B and Table I). The masses of the proteolytic fragments were determined with sufficient accuracy to establish that the 26,611-Da product corresponds to the amino acid stretch Thr-201 to Glu-432 (calculated mass, 26,651 Da) (Fig. 6A), whereas the 26,424-Da fragment can be assigned to the fragment Gln-203 to Glu-432 (calculated mass, 26,402 Da) (Fig. 6B). Moreover, with both elastase and supernatant, the initial fragment VE-EC1-4 (calculated mass, 48,941 Da) could still be detected (Fig. 6, A and C).


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Fig. 6.   Identification of elastase by MALDI mass spectrometry as the only protease released into the cell culture medium following adhesion of neutrophils on endothelial monolayers. Supernatants used in this figure were obtained in conditions identical to those depicted in Fig. 3 (lane 6). VE-EC1-4 (3 µM) was mixed with purified elastase (0.02 unit/ml, panel A), with purified cathepsin G (0.02 unit/ml, panel B), or with supernatant (dilution 1/3, panel C) and incubated at 37 °C during 2 h. Following proteolysis, the various mixtures were analyzed by MALDI mass spectrometry. In panel B, based on band intensity variations, the 19,666- and 16,476-Da products were probably generated from the 22,606-Da fragment. D, prior to the addition to VE-EC1-4, the diluted supernatant (1/3) was mixed with purified cathepsin G (0.02 unit/ml). Incubation and mass spectrometry were performed as previously described in panels A-C. The inset presents an enlargement of the mass spectrum corresponding to VE-EC1-4 digestions either by the supernatant alone (dotted line) or by the cathepsin G-treated supernatant (continuous line). The arrow indicates an additional proteolytic product resulting from the addition of cathepsin G to the supernatant. The asterisk marks an artifact resulting from the use of matrix-containing sinapinic acid.

The 38-kDa fragments (Figs. 4 and 5) were not detected by mass spectrometry. This was probably a result of the method used for treating the samples before mass spectrometry analysis. Indeed, to increase their concentration, samples were passed through a ZipTip column. Analysis by Western blot of the samples prior to and after elution on ZipTip columns indicated that large amounts of the 38-kDa fragments remained fixed on the columns. This explains the discrepancy of protein fragment quantities observed between Western blot and mass spectrometry analyses. In contrast, MALDI mass spectrometry analyses detected additional fragments. These fragments generated by elastase or cathepsin G, respectively, possess molecular masses of 22,300 Da (Fig. 6A) or 22,669 Da (Fig. 6B) and correspond to the N-terminal peptide fragments MD1-V200 (calculated mass, 22,300 Da) or MD1-Q203 (calculated mass, 22,669 Da). These bands are absent from the Western blots revealed with the antibody anti-Cad3 (Figs. 4 and 5) but are detected with the antibody anti-EC1 as previously mentioned.

Altogether, mass spectrometry and N-terminal sequencing analyses indicate that elastase cleaves the VE-EC1-4 fragment after residues Ile-93 and Val-200, whereas cathepsin G cleaves after residues Phe-91, Lys-94, and Gln-203, as illustrated in Table I.

MALDI mass spectrometry combined to N-terminal sequencing allowed us to clearly identify the nature of the protease secreted by neutrophils following their adhesion on endothelial cell monolayers. Indeed, the supernatant contained a protease that generated a 26,625-Da fragment, similarly to elastase but different from the 26,424-Da cathepsin G-generated fragment (Fig. 6C). This result strongly suggests that elastase is probably the only protease released from neutrophils that is capable to digest VE cadherin. To verify this hypothesis, we added purified cathepsin G to the supernatant prior to its addition to the VE-EC1-4 fragment (Fig. 6D). MALDI analysis exhibited two major cleavage products of 26,604 and 26,395 Da, corresponding, as previously established, to fragments Thr-201 to Glu-432 and Gln-203 to Glu-432, respectively. This result indicated that, if cathepsin G would have been released into the culture medium, it could easily be detected by mass spectrometry.

The selective secretion of elastase in the supernatant ruled out the possibility that VE cadherin cleavage resulted from neutrophil crushing during the cell purification step. In this case, both elastase and cathepsin G would have been detected in cell supernatants.

Cleavage of the Extracellular Domain of VE Cadherin by Elastase and Cathepsin G Bound to the Leukocyte Cell Surface-- Several papers mention that exposure of neutrophils to cytokines or chemoattractants such as fMLP and TNFalpha induces translocation of elastase and cathepsin G from the azurophil granules to the external surface plasma membrane of neutrophils (36, 38, 39). These surface-bound proteases were demonstrated to retain their enzymatic activity.

It thus remains to be shown whether, following adhesion of neutrophils on endothelial cell monolayers, the cleavage of VE cadherin is induced by proteases secreted into the extracellular milieu or bound to the neutrophil surface membrane. To answer this question, neutrophils, stimulated both by fMLP and TNFalpha , were first fixed with paraformaldehyde and glutaraldehyde and subsequently incubated with the VE-EC1-4 fragment. Western blot analysis showed that the resulting digestion pattern was comparable with that previously observed using purified elastase or cathepsin G (Fig. 7A, lanes 2, 5, and 8). Indeed, the typical fragments of 38 and 27 kDa generated by leukocyte enzymes were clearly detected, indicating the presence of active elastase or cathepsin G at the cell surface of neutrophils.


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Fig. 7.   Proteolysis of VE-EC1-4 by elastase and cathepsin G bound to the leukocyte cell surface. A, Western blot analysis of VE-EC1-4 proteolysis. Neutrophils were incubated with 10-8 M fMLP and 100 units/ml TNFalpha for 30 min and then fixed. Fragment VE-EC1-4 (5 µg) was incubated for 3 h at 37 °C with purified elastase (7 × 10-4 units) (lanes 2-4), purified cathepsin G (7 × 10-4 units) (lanes 5-7), or fixed neutrophils (105 cells) (lane 8). The leukocyte protease inhibitor SLPI (10-10 mol) (lanes 3 and 6) or the elastase-specific inhibitor MeOSuc-AAPV (10-10 mol) (lanes 4 and 7) were added. Lane 1 corresponds to VE-EC1-4 alone. Note the 38- and 27-kDa immunoblotted bands in the presence of fixed neutrophils. B, MALDI mass spectrometry analysis of the VE-EC1-4-derived proteolytic products. Mixtures containing both fixed neutrophils and VE-EC1-4 were centrifuged and the supernatants collected. MALDI analysis performed on these supernatants showed a double peak (*) representing proteolytic fragments with molecular masses of 27 kDa and a 49-kDa peak corresponding to residual uncleaved VE-EC1-4 (**). The inset shows an enlargement of the 27-kDa doublet (*).

This Western blot analysis did not allow to discriminate whether bound elastase or bound cathepsin G or both were involved in the cleavage of VE cadherin. In contrast, MALDI mass spectrometry analysis revealed two distinct major products having molecular masses of 26,615 and 26,395 Da (Fig. 7B). They were identical to those generated by elastase and by cathepsin G as previously demonstrated. To exclude the possibility that this proteolysis was caused by the release of enzymes from intracellular granules, postwashing supernatants from fixed neutrophils were also incubated with VE-EC1-4. In these conditions, no proteolytic product was detected by mass spectrometry (data not shown). This indicated that the proteolytic activity was cell-associated and not a result of the release of intracellular leukocyte-derived enzymes. Altogether, data indicated that both elastase and cathepsin G bound to neutrophil cell membranes could cleave the extracellular part of VE cadherin.

Elastase- and Cathepsin G-mediated Increase of Endothelial Monolayer Permeability-- To test whether elastase or cathepsin G could be responsible for the opening of inter-endothelial cell junctions during transmigration of leukocytes, transendothelial permeability was measured in the presence of these purified proteases. The modification of permeability was quantified by establishing the transmigration time course of a peroxidase-conjugated anti-mouse IgG across endothelial cell monolayers seeded on porous Transwell chambers (Fig. 8). A clear accumulation of the marker with time was observed in the lower compartments for elastase-treated monolayers compared with untreated ones (Fig. 8A). Similar results were observed when elastase was replaced by either cathepsin G (Fig. 8B) or by supernatants collected from endothelial cells on which neutrophils had adhered (Fig. 8C). This last observation was consistent with the fact that these cell supernatants, as previously demonstrated, contained elastase. In fact, permeability increased with time and reached a plateau 90 min after the addition of proteases. It can be deduced that resealing of adherent junctions takes place after 90 min of incubation, thus preventing additional accumulation of the marker in the lower compartment. This confirmed our previous finding obtained with an anti-VE cadherin antibody, which was able to transiently increase transendothelial permeability by destabilizing endothelial cell-cell junctions (18). Altogether, these results suggest both elastase and cathepsin G are able to perturb the integrity of the endothelial cell monolayer likely by damaging VE cadherin at cell-cell junctions.


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Fig. 8.   Time course of transendothelial permeability induced by either elastase or cathepsin G or cell supernatants. Supernatants used in this figure were obtained in conditions identical to those depicted in Fig. 3 (lane 6). Cell permeability was quantified by measuring the transmigration of peroxidase-conjugated anti-mouse IgG across endothelial cell monolayers seeded on porous Transwell filters. Practically, once cells reached confluence, mixtures of either elastase (2 × 10-2 units/ml) and marker (panel A, curve 1) or cathepsin G (2 × 10-2 units/ml) and marker (panel B, curve 1) or supernatant and marker (panel C, curve 1) were added to the upper compartments of Transwell units. In the controls (curves 2), only the marker was added. Optical density represents the amount of marker accumulated for the different incubation times in the lower compartments. They correspond to mean values obtained from three similarly treated independent Transwell units.

Neutrophil Transmigration Blockage by Protease Inhibitors-- To assess the role of elastase and cathepsin G in neutrophil transmigration, an in vitro assay was performed using Transwell units. Differentially treated neutrophils were added to confluent endothelial cell monolayers seeded on porous membranes and pre-activated with TNFalpha . Prior to their addition to endothelial cell monolayers, freshly purified neutrophils underwent specific treatments. First, they were treated with both TNFalpha and fMLP in such a way that maximal amounts of elastase and cathepsin G were translocated at the neutrophil external membrane (36); second, to block the surface-bound proteases, TNFalpha - and fMLP-stimulated neutrophils were incubated with either SLPI or MeOSuc-AAPV. This procedure was necessary to avoid direct contact of SLPI with endothelial cells, an event that increases by itself the permeability of the endothelial monolayers (data not shown).

To detect the presence or the absence of protease activity at the neutrophil cell surface, differentially treated neutrophils were fixed before being incubated with fragment VE-EC1-4. The typical fragments of 38 and 27 kDa previously described in Fig. 7 are detected when VE-EC1-4 was incubated with untreated neutrophils attesting that some proteases are bound to the surface of these neutrophils (Fig. 9A, lane 2). Treatment of neutrophils with both TNFalpha and fMLP increases the amount of these proteolytic fragments slightly (Fig. 9A, lane 3). By contrast, they disappear when VE-EC1-4 was incubated with either SLPI- or MeOSuc-AAPV-treated neutrophils (Fig. 9A, lanes 4 and 5). This indicates that the addition of protease inhibitors fully hampers protease activity at the cell surface of TNFalpha - and fMLP-stimulated neutrophils. However, a subsequent 30-min incubation of the SLPI-treated neutrophils with TNFalpha induced a new translocation of proteases at the neutrophil cell surface. Indeed, after this second contact with TNFalpha , the 38- and 27-kDa fragments reappeared as illustrated in Fig. 9B (lanes 1-3).


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Fig. 9.   Inhibition of neutrophil transmigration by protease inhibitors. A, inhibition of proteases at the neutrophil cell surface. The neutrophils were stimulated with both TNFalpha (100 units/ml) and fMLP (10-8 M) during 30 min at 37 °C. They were then rinsed twice with PBS and incubated with SLPI (2.5 × 10-10 mol) (lane 4), MeOSuc-AAPV (2 × 10-6 mol) (lane 5), or PBS (lane 3) for an additional 2 h. TNFalpha - and fMLP-stimulated (lanes 3-5) or non-stimulated (lane 2) neutrophils (105 cells) were fixed with paraformaldehyde and glutaraldehyde prior to incubation with VE-EC1-4 (5 µg) for 30 min. For comparison, undigested VE-EC1-4 is shown (lane 1). Western blot analysis was then performed on the digested products as described in Fig. 3. In the presence of SLPI or MeOSuc-AAPV, the 38- and 27-kDa proteolytic products disappeared. B, stimulation of cell surface expression of neutrophil proteases. TNFalpha - and fMLP-stimulated neutrophils were treated (lane 2) or not (lane 1) with SLPI as previously described in panel A of this figure. TNFalpha - and fMLP-stimulated neutrophils that were also treated with SLPI (lane 2) were stimulated a second time with TNFalpha and fMLP during 30 min (lane 3). The differently treated neutrophils were then fixed as previously described prior to incubation with VE-EC1-4 for 30 min. Western blot analysis was then performed on the digested products as described in Fig. 3. The second TNFalpha and fMLP stimulation induced the apparition of the 38- and 27-kDa proteolytic products. C, transmigration assay of neutrophils. TNFalpha - and fMLP- stimulated neutrophils treated with either SLPI (lane 3) or MeOSuc-AAPV (lane 4) or untreated with inhibitor (lane 2) were added to the upper compartments of the Transwell units at 5 × 104 cells/insert. For comparison, non-stimulated neutrophils were added to inactivated endothelial cell monolayers (lane 1). 30 min after the addition of neutrophils, the lower compartments of the Transwell units were centrifuged and the supernatants discarded. Extraction and quantification of DNA from the transmigrated neutrophils enabled us to count their number. Solid bars represent the means of four experiments, and their calculated errors are shown.

TNFalpha - and fMLP-stimulated neutrophils easily transmigrate across TNFalpha -activated endothelial cell monolayers (Fig. 9C, lane 2). Inhibition of proteases by SLPI or MeOSuc-AAPV significantly reduces the neutrophil transmigration rate (Fig. 9C, lanes 3 and 4). The partial inhibition of transmigration can be the result of reappearance of proteases at the neutrophil cell surface following their contact with TNFalpha -activated endothelial cell monolayers. Furthermore, some untreated neutrophils transmigrate across inactivated endothelial cells (Fig. 9C, lane 1) probably attracted by the presence of fMLP in the lower compartments of Transwell units. These results indicate that elastase and cathepsin G localized at the neutrophil cell surface participate in the transmigration process probably by locally cleaving VE cadherin.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The mechanism by which neutrophils transmigrate across the vascular endothelium is controversial. Some studies suggest that adherent neutrophils trigger intracellular events in the endothelium, which lead to the disorganization of the junctional complexes involved in the maintenance of endothelial integrity (4, 21, 22). Other studies indicate that these junctional complexes disrupt under the action of neutrophil-derived proteases, thus facilitating neutrophil migration (30, 34).

Herein, we show that VE cadherin, an adhesive receptor participating at the maintenance of endothelial integrity (18), is cleaved following adhesion of neutrophils to endothelial cell monolayers.

Indeed, small but detectable amounts of soluble fragments of VE cadherin accumulated in the cell culture supernatants after adhesion of neutrophils. We can exclude an artifactual degradation of VE cadherin by released proteases caused by neutrophil lysis in our experiments. Therefore, the low level of soluble VE cadherin fragments probably reflects a physiologically relevant, local proteolysis activity close to the very sites of neutrophil adhesion.

Using specific inhibitors of neutrophil proteases, we were able to identify elastase and cathepsin G as the major proteases involved in the cleavage of VE cadherin. There is a redundancy between these two proteases to cleave VE cadherin. To abolish neutrophil transmigration, these cells must at least be deficient in both elastase and cathepsin G. This can explain why neutrophils still transmigrate across endothelial monolayers in the presence of specific inhibitors of elastase and also why neutrophils from elastase-deficient mice show no defect in transendothelial migration (33). Indeed, cathepsin G can replace elastase, thus allowing the neutrophil transmigration to occur despite elastase inhibition or deficiency. Redundancy is a commonly observed feature of complex biological processes allowing for flexibility under different physiological conditions.

Using MALDI-time of flight mass spectrometry and N-terminal sequencing, we demonstrate that purified elastase can cleave the recombinant fragment VE-EC1-4 overlapping the four-N-terminal extracellular modules of VE cadherin after the amino acids Ile-93 and Val-200. Similarly, purified cathepsin G produces three cleavage sites on VE-EC1-4 located after amino acids Phe-91, Lys-94, and Gln-203, i.e. shifted by only two or three amino acids when compared with those of elastase. All of these cleavage sites map within the EC1-EC2 and the EC2-EC3 interdomain regions of VE cadherin, which are structurally more prone to be exposed than the compact domains they connect. The recombinant fragment VE-EC1-5, extended by the C-terminal module EC5 and corresponding to the complete extracellular domain of VE cadherin, was also used in these proteolysis experiments (data not shown). Because of the heterogeneity of its glycosylation, it cannot be used for determining the position of the cleavage sites. Nevertheless, when digested by elastase and cathepsin G, its digestion pattern appears more complex compared with that of VE-EC1-4. This suggests that these two proteases possess additional cleavage sites localized between the ends of modules EC4 and EC5 of VE cadherin.

Both fragments VE-EC1-4 and VE-EC1-5 elaborate a hexameric structure in solution (12),2 probably reflecting the self-association of VE cadherin at the surface of endothelial cells. Based on our previous studies (11, 12), we know that the extracellular module EC1 and the intact EC3-EC4 tandem-module (amino acid stretch T212-E431) are both required for VE cadherin self-assembly. Consequently, at the endothelial cell surface, elastase and cathepsin G may cleave the VE cadherin molecule at positions identical to those observed for the recombinant fragment VE-EC1-5, thus probably destroying its hexameric assembly. Based on our previous knowledge on the self-assembly of VE cadherin, the short fragments left at the endothelial cell surface membrane are probably unable to self associate. We know that destabilization of VE cadherin homophilic interactions by an anti-VE cadherin antibody alters the integrity of endothelial cell monolayers by creating numerous gaps at cell-cell junctions (18). Similarly, cleavage of VE cadherin by elastase or cathepsin G could induce formation of gaps between endothelial cells through which neutrophils could migrate from the vasculature into the underlying tissues.

We demonstrate that adhesion of neutrophils on endothelial cell monolayers induces the release of elastase from neutrophils into the supernatant of the cell culture. In contrast, no cathepsin G was extracellularly detected, although its intracellular content within azurophil granules is as abundant as that of elastase (38). Because of a different surface charge, elastase does not remain bound to the neutrophil outer membrane and therefore is released into the extracellular milieu (36, 38-40). Our results also confirm that, following neutrophil stimulation by TNFalpha and fMLP, elastase and cathepsin G are present at the cell surface whereas they are mainly stored within internal azurophil granules in resting cells (36). Our data indicate that both cell surface-bound enzymes are active and sufficiently accessible to be able to cleave, similarly to their soluble forms, the VE-EC1-4 fragment. This result suggests that, following close contact between neutrophils and endothelial cells, cell surface proteases may locally cleave VE cadherin.

Furthermore, we prove that purified elastase and cathepsin G are able to increase endothelial monolayer permeability in vivo using Transwell units. It can therefore be deduced that treatment of endothelial cell monolayers with elastase or cathepsin G disrupt cell-cell junctions, probably by proteolysing VE cadherin molecules involved in endothelium monolayer integrity.

Altogether, our data suggest that neutrophil transmigration is facilitated by elastase secreted into the extracellular milieu and by membrane-bound elastase or cathepsin G despite specific inhibitors such as alpha 1-antitrypsin or SLPI within the blood. Indeed, in zones of close contact between neutrophils and endothelial cells, the concentration of soluble elastase is several orders of magnitude higher than the serum concentration of inhibitors. Consequently, the soluble enzymes may transiently overwhelm the local inhibitor concentration, thus allowing VE cadherin proteolysis to take place (41). Moreover, it is possible that membrane-bound elastase and cathepsin G may be more resistant to inhibition when compared with secreted elastase, inhibitors being unable to reach the zones of tight adhesion between neutrophils and endothelial cells (31). The action of these physiological inhibitors confines the activity of elastase and cathepsin G to the very sites of neutrophil adhesion on endothelial cells and thus limits their damaging activity.

The leukocyte transmigration through the endothelium is a multistep process. First, the leukocyte must firmly adhere to the endothelium involving dynamic interactions between the leukocyte integrin alpha Lbeta 2 and the endothelial immunoglobulin superfamily proteins such as JAM-1 (42) and ICAM-1. Homophilic interactions mediated by PECAM-1 (43) and CD99 (7) act in concert to facilitate the progression of the leukocyte across the adherens junctions. During this transendothelial migration, VE cadherin moves away to different ends of the transmigration site (4, 29). The event initiating the lateral movement of VE cadherin may correspond to the cleavage of this adhesive receptor by surface-bound elastase or/and cathepsin G. The subsequent disruption of its hexameric assembly might open the way at the front of leukocyte migration.

    ACKNOWLEDGEMENTS

We are indebted to staff of Hôpital Sud (Grenoble, France) for kindly collecting umbilical cords for these experiments.

    FOOTNOTES

* This work was supported in part by Grant 4447 from the Association pour la Recherche sur le Cancer and from Groupement des Entreprises Françaises dans la Lutte contre le Cancer.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.

§ Recipient of a fellowship from Association de Recherche sur la Polyarthrite.

|| Current address: European Molecular Biology Laboratory, 38 042, Grenoble, France.

** To whom correspondence should be addressed. E-mail: gulino@ibs.fr.

Published, JBC Papers in Press, February 12, 2003, DOI 10.1074/jbc.M300351200

2 S. Bibert, E. Concord, E. Hewat, B. Dublet, T. Vernet, and D. Gulino-Debrac, manuscript in preparation.

    ABBREVIATIONS

The abbreviations used are: PECAM, platelet-endothelial cell adhesion molecule; VE cadherin, vascular endothelium cadherin; CAPS, 3-(cyclohexylamino)-1-propanesulfonic acid; ELISA, enzyme-linked immunosorbent assay; HUVEC, human umbilical vein endothelial cell; fMLP, formyl-Met-Leu-Phe; MALDI, matrix-assisted laser desorption; SLPI, secretory leukocyte protease inhibitor; TNFalpha , tumor necrosis factor alpha ; MeOSuc-AAPV, N-methoxysuccinyl-Ala-Ala-Pro-Val chloromethyl ketone; PBS, phosphate-buffered saline.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1. Worthylake, R. A., and Burridge, K. (2001) Curr. Opin. Cell Biol. 13, 569-577[CrossRef][Medline] [Order article via Infotrieve]
2. Dunon, D., Piali, L., and Imhof, B. A. (1996) Curr. Opin. Cell Biol. 8, 714-723[CrossRef][Medline] [Order article via Infotrieve]
3. Johnson-Leger, C., Aurrand-Lions, M., and Imhof, B. A. (2000) J. Cell Sci. 113, 921-933[Abstract/Free Full Text]
4. Shaw, S. K., Bamba, P. S., Perkins, B. N., and Luscinskas, F. W. (2001) J. Immunol. 167, 2323-2330[Abstract/Free Full Text]
5. Feng, D., Nagy, J. A., Pyne, K., Dvorak, H. F., and Dvorak, A. M. (1998) J. Exp. Med. 187, 903-915[Abstract/Free Full Text]
6. Newman, P. J. (1999) J. Clin. Invest. 103, 5-9[Free Full Text]
7. Schenkel, A. R., Mamdouh, Z., Chen, X., Liebman, R. M., and Muller, W. A. (2002) Nat. Immunol. 3, 143-150[CrossRef][Medline] [Order article via Infotrieve]
8. Rival, Y., Del Maschio, A., Rabiet, M. J., Dejana, E., and Duperray, A. (1996) J. Immunol. 157, 1233-1241[Abstract]
9. Vaporciyan, A. A., DeLisser, H. M., Yan, H. C., Mendiguren, I. I., Thom, S. R., Jones, M. L., Ward, P. A., and Albelda, S. M. (1993) Science 262, 1580-1582[Medline] [Order article via Infotrieve]
10. Lampugnani, M. G., Resnati, M., Raiteri, M., Pigott, R., Pisacane, A., Houen, G., Ruco, L. P., and Dejana, E. (1992) J. Cell Biol. 118, 1511-1522[Abstract]
11. Bibert, S., Jaquinod, M., Concord, E., Ebel, C., Hewat, E., Vanbelle, C., Legrand, P., Weidenhaupt, M., Vernet, T., and Gulino-Debrac, D. (2002) J. Biol. Chem. 277, 12790-12801[Abstract/Free Full Text]
12. Legrand, P., Bibert, S., Jaquinod, M., Ebel, C., Hewat, E., Vincent, F., Vanbelle, C., Concord, E., Vernet, T., and Gulino, D. (2001) J. Biol. Chem. 276, 3581-3588[Abstract/Free Full Text]
13. Ozawa, M., Baribault, H., and Kemler, R. (1989) EMBO J. 8, 1711-1717[Abstract]
14. Butz, S., Stappert, J., Weissig, H., and Kemler, R. (1992) Science 257, 1142-1144[Medline] [Order article via Infotrieve]
15. Jou, T. S., Stewart, D. B., Stappert, J., Nelson, W. J., and Marrs, J. A. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 5067-5071[Abstract]
16. Rimm, D. L., Koslov, E. R., Kebriaei, P., Cianci, C. D., and Morrow, J. S. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 8813-8817[Abstract]
17. Ferber, A., Yaen, C., Sarmiento, E., and Martinez, J. (2002) Exp. Cell Res. 274, 35-44[CrossRef][Medline] [Order article via Infotrieve]
18. Gulino, D., Delachanal, E., Concord, E., Genoux, Y., Morand, B., Valiron, M. O., Sulpice, E., Scaife, R., Alemany, M., and Vernet, T. (1998) J. Biol. Chem. 273, 29786-29793[Abstract/Free Full Text]
19. Corada, M., Mariotti, M., Thurston, G., Smith, K., Kunkel, R., Brockhaus, M., Lampugnani, M. G., Martin-Padura, I., Stoppacciaro, A., Ruco, L., McDonald, D. M., Ward, P. A., and Dejana, E. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 9815-9820[Abstract/Free Full Text]
20. Hordijk, P. L., Anthony, E., Mul, F. P., Rientsma, R., Oomen, L. C., and Roos, D. (1999) J. Cell Sci. 112 12, 1915-1923
21. Allport, J. R., Ding, H., Collins, T., Gerritsen, M. E., and Luscinskas, F. W. (1997) J. Exp. Med. 186, 517-527[Abstract/Free Full Text]
22. Del Maschio, A., Zanetti, A., Corada, M., Rival, Y., Ruco, L., Lampugnani, M. G., and Dejana, E. (1996) J. Cell Biol. 135, 497-510[Abstract]
23. Rabiet, M. J., Plantier, J. L., Rival, Y., Genoux, Y., Lampugnani, M. G., and Dejana, E. (1996) Arterioscler. Thromb. Vasc. Biol. 16, 488-496[Abstract/Free Full Text]
24. Carmeliet, P. (2000) Nat. Med. 6, 389-395[CrossRef][Medline] [Order article via Infotrieve]
25. Shay-Salit, A., Shushy, M., Wolfovitz, E., Yahav, H., Breviario, F., Dejana, E., and Resnick, N. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 9462-9467[Abstract/Free Full Text]
26. Nawroth, R., Poell, G., Ranft, A., Kloep, S., Samulowitz, U., Fachinger, G., Golding, M., Shima, D. T., Deutsch, U., and Vestweber, D. (2002) EMBO J. 21, 4885-4895[Abstract/Free Full Text]
27. Ukropec, J. A., Hollinger, M. K., Salva, S. M., and Woolkalis, M. J. (2000) J. Biol. Chem. 275, 5983-5986[Abstract/Free Full Text]
28. Ukropec, J. A., Hollinger, M. K., and Woolkalis, M. J. (2002) Exp. Cell Res. 273, 240-247[CrossRef][Medline] [Order article via Infotrieve]
29. Su, W. H., Chen, H. I., and Jen, C. J. (2002) Blood 100, 3597-3603[Abstract/Free Full Text]
30. Cepinskas, G., Sandig, M., and Kvietys, P. R. (1999) J. Cell Sci. 112, 1937-1945[Abstract/Free Full Text]
31. Owen, C. A., and Campbell, E. J. (1995) Semin. Cell Biol. 6, 367-376[Medline] [Order article via Infotrieve]
32. Moll, T., Dejana, E., and Vestweber, D. (1998) J. Cell Biol. 140, 403-407[Abstract/Free Full Text]
33. Allport, J. R., Lim, Y. C., Shipley, J. M., Senior, R. M., Shapiro, S. D., Matsuyoshi, N., Vestweber, D., and Luscinskas, F. W. (2002) J. Leukocyte Biol. 71, 821-828[Abstract/Free Full Text]
34. Carden, D., Xiao, F., Moak, C., Willis, B. H., Robinson-Jackson, S., and Alexander, S. (1998) Am. J. Physiol. 275, H385-H392[Medline] [Order article via Infotrieve]
35. Weiss, J., Kao, L., Victor, M., and Elsbach, P. (1985) J. Clin. Invest. 76, 206-212[Medline] [Order article via Infotrieve]
36. Owen, C. A., Campbell, M. A., Sannes, P. L., Boukedes, S. S., and Campbell, E. J. (1995) J. Cell Biol. 131, 775-789[Abstract]
37. Weinrauch, Y., Drugan, D., Shapiro, S. D., Weiss, J., and Zychlinsky, A. (2002) Nature 417, 91-94[CrossRef][Medline] [Order article via Infotrieve]
38. Campbell, E. J., Campbell, M. A., and Owen, C. A. (2000) J. Immunol. 165, 3366-3374[Abstract/Free Full Text]
39. Owen, C. A., Campbell, M. A., Boukedes, S. S., and Campbell, E. J. (1995) J. Immunol. 155, 5803-5810[Abstract]
40. Owen, C. A., Campbell, M. A., Boukedes, S. S., and Campbell, E. J. (1997) Am. J. Physiol. 272, L385-L393[Medline] [Order article via Infotrieve]
41. Stockley, R. A. (2001) Novartis Found. Symp. Disc. 234, 189-204
42. Ostermann, G., Weber, K. S., Zernecke, A., Schroder, A., and Weber, C. (2002) Nat. Immunol. 3, 151-158[CrossRef][Medline] [Order article via Infotrieve]
43. Vestweber, D. (2002) Curr. Opin. Cell Biol. 14, 587-593[CrossRef][Medline] [Order article via Infotrieve]
44. Gotsch, U., Borges, E., Bosse, R., Boggemeyer, E., Simon, M., Mossmann, H., and Vestweber, D. (1997) J. Cell Sci. 110, 583-588[Abstract/Free Full Text]


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