Potential role for mast cell tryptase in recruitment of inflammatory cells to endothelium

Maureen C. Meyer, Michael H. Creer, and Jane McHowat

Department of Pathology, Saint Louis University School of Medicine, St. Louis, Missouri

Submitted 4 May 2005 ; accepted in final form 28 July 2005


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Recent research suggests that activation of protease-activated receptors (PARs) on the surface of endothelial and epithelial cells may play a role in general mechanisms of inflammation. We hypothesized that mast cell tryptase activation of endothelial cell PAR-2 is coupled to increased calcium-independent PLA2 (iPLA2) activity and increased platelet-activating factor (PAF) production that may play a role in inflammatory cell recruitment at sites of vascular injury. Stimulation of human coronary artery endothelial cells (HCAEC) with 20 ng/ml tryptase increased iPLA2 activity, arachidonic acid release, and PAF production. These tryptase-stimulated responses were inhibited by pretreatment with the iPLA2-selective inhibitor bromoenol lactone (BEL; 5 µM, 10 min). Similar patterns of increased iPLA2 activity and PAF production were also seen when HCAEC were treated with SLIGKV, which represents the tethered ligand sequence for the human PAR-2 once the receptor is cleaved by tryptase. Tryptase stimulation also increased cell surface expression of P-selectin, decreased electrical resistance, and increased neutrophil adherence to the endothelial cell monolayer. The tryptase-stimulated increases in both cell surface P-selectin expression and neutrophil adhesion were also inhibited with BEL pretreatment. We conclude that tryptase stimulation of HCAEC contributes importantly to early inflammatory events after vascular injury by activation of iPLA2, leading to arachidonic acid release, PAF production, cell surface P-selectin expression, and increased neutrophil adherence.

atherosclerosis; endothelial cells


THE PROTEASE-ACTIVATED RECEPTOR (PAR) represents a family with four currently recognized forms (PAR-1 to PAR-4) that are activated by a unique mechanism involving proteolytic cleavage of the receptor amino terminus and are coupled to G proteins (see Ref. 5 for review). Evidence suggests that interaction of proteases with PAR has far-reaching implications in diversified cellular responses, particularly in general mechanisms of inflammation and host defense (2, 26). PAR may play important roles in both the acute and chronic inflammatory responses of both endothelial and epithelial cells that form the defensive barriers of the body (2). Activation of the endothelium through the activation of PAR-1 by thrombin has been shown to result in activation of phospholipases, enzymes known to play an essential role in inflammation (4). PARs couple to multiple intracellular signaling pathways that are related to growth and inflammation, including activation of phospholipases, protein kinase C, and MAP kinases (4).

The presence of PAR-1 and PAR-2 on the endothelial cell (EC) surface extends the number of proteases to which the cells respond rather than being coupled to different intracellular responses (22). Intracellular signaling events coupled to PAR-1, which is activated by thrombin, have been, and continue to be, the most thoroughly studied. PAR-2 receptors are activated by tryptase, a protease that is produced only by mast cells (8). Mast cells are virtually always found in close spatial proximity to EC, which provide stem cell factor (SCF or c-Kit ligand) required for mast cell survival (SCF actively suppresses apoptosis in mast cells; Ref. 13). In particular, studies demonstrate that mast cells are abundant in thrombosed veins and in coronary atheromas (14). Although PAR-2 activation by mast cell tryptase may play an important role in EC responses in these settings, intracellular signaling events after PAR-2 activation have not been studied as intensively as PAR-1.

Accordingly, this study investigated whether tryptase stimulation, in the absence or presence of the calcium-independent PLA2 (iPLA2)-selective inhibitor bromoenol lactone (BEL), of PAR-2 on human coronary artery endothelial cells (HCAEC) activates iPLA2 and leads to increased platelet-activating factor (PAF) production and arachidonic acid release. Because PAF was previously shown to contribute importantly to EC recruitment of circulating inflammatory cells by a process that begins with P-selectin-mediated binding of leukocytes (polymorphonuclear neutrophils, PMN) (23), we also determined whether mast cell tryptase activation of HCAEC PAR-2 would contribute to inflammatory cell recruitment via upregulation of P-selectin expression and enhanced PMN binding to tryptase-stimulated EC. On activation, mast cells release numerous factors including histamine, proteases, chemotactic factors, cytokines, and metabolites of arachidonic acid, many capable of modulating adhesion molecule expression on the EC surface. To examine specifically the role of tryptase in stimulating the endothelium, we choose to treat HCAEC with tryptase rather than culturing them with activated mast cells.


    METHODS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents. Human tryptase (200 µg/ml recombinant skin {beta}-tryptase with 0.5 mg/ml heparin) was purchased from Promega (Madison, WI). BEL was obtained from Cayman Chemical (Ann Arbor, MI). Goat anti-P-selectin antibody and horseradish peroxidase-conjugated rabbit anti-goat antibody were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). [3H]Arachidonic acid and [3H]acetic acid were obtained from Perkin Elmer Life Sciences (Boston, MA). All other reagents were purchased from Sigma Chemical (St. Louis, MO).

Culture of EC. HCAEC were obtained from Cambrex (Walkersville, MD). Cells were grown to 80–90% confluence in EGM-2MV medium (Cambrex) and passaged with the Subculture Reagent Pack (Cambrex). For experiments, ~3 x 105 cells in 2 ml of EGM-2MV medium were placed in 34-mm culture dishes (Corning, Corning, NY). Unless otherwise stated, cells from passages 3–4 were used for experiments.

Immunoblot analysis. HCAEC were removed from the tissue culture plate in lysis buffer containing (in mM) 20 HEPES, 250 sucrose, 2 DTT, 2 EDTA, 2 EGTA, 10 {beta}-glycerophosphate, 1 sodium orthovanadate, and 2 phenylmethylsulfonyl fluoride, with 20 µg/ml leupeptin, 10 µg/ml aprotinin, and 5 µg/ml pepstatin A (pH 7.6). Protein was mixed with an equal volume of SDS sample buffer and heated at 95°C for 5 min before loading onto a 10% polyacrylamide gel. Protein was separated by SDS-PAGE at 200 V for 40 min and electrophoretically transferred to polyvinylidene difluoride (PVDF-Plus) membranes (Micron Separations, Westborough, MA) at 100 V for 1 h. Nonspecific sites were blocked by incubating the membranes with Tris-buffered saline containing 0.1% (vol/vol) Tween 20 (TBST) and 5% (wt/vol) nonfat milk for 1 h at room temperature. The blocked PVDF membrane was incubated with primary antibody to PAR-2 for 2 h at room temperature. Unbound antibody was removed with five washes with TBST, and membrane was incubated with horseradish peroxidase-conjugated secondary antibody. After five washes with TBST, regions of antibody binding were detected with enhanced chemiluminescence (SuperSignal; Pierce, Rockford, IL) and exposure to film (Hyperfilm; Amersham).

Tryptase stimulation. Human recombinant skin {beta}-tryptase (Promega) was diluted with medium (iPLA2 assay, arachidonic acid release, resistance measurements, and neutrophil adhesion assay), or Hanks' balanced salt solution (PAF production and P-selectin surface expression assays) to the working concentration. Tryptase was added to the cell culture plate, and the plate was gently rotated to ensure thorough mixing and even distribution of tryptase across the HCAEC monolayer.

SLIGKV stimulation. SLIGKV (Invitrogen, Carlsbad, CA) was diluted with medium (iPLA2 assay) or Hanks' balanced salt solution (PAF production) to the working concentration. Cells were stimulated with 100 µM SLIGKV. SLIGKV was added to the cell culture plate, and the plate was gently rotated to ensure thorough mixing and even distribution of SLIGKV across the HCAEC monolayer.

Measurement of PLA2 activity. HCAEC were grown to confluence and stimulated with 20 ng/ml tryptase for 0–60 min. At the end of the stimulation period, the medium was removed from the HCAEC monolayer and replaced with ice-cold PLA2 buffer containing (in mM) 250 sucrose, 10 KCl, 10 imidazole, 5 EDTA, and 2 DTT, with 10% glycerol (pH 7.8). The cells were removed from the tissue culture dish by scraping, and the suspension was sonicated on ice for three bursts of 10 s.

PLA2 activity in the cell sonicates was assessed by incubating enzyme (50 µg of protein) with 100 µM (16:0, [3H]18:1) plasmenylcholine substrate (synthesized as described in Ref. 21) in assay buffer containing 10 mM Tris, 10% glycerol, and 4 mM EGTA, pH 7.0, at 37°C for 5 min in a total volume of 200 µl. Reactions were initiated by adding the radiolabeled phospholipid substrate as a concentrated stock solution in ethanol and terminated by the addition of 100 µl of butanol. Released radiolabeled fatty acid was isolated and quantified by application of 25 µl of the butanol phase to channeled Silica Gel G plates, development in petroleum ether-diethyl ether-acetic acid (70:30:1, vol/vol/vol), and liquid scintillation spectrometry.

Arachidonic acid release. The extent of arachidonic acid release was determined by measuring the amount of [3H]arachidonic acid released into the surrounding medium from HCAEC prelabeled with 3 µCi of [3H]arachidonic acid per 34-mm culture dish (Corning) for 18 h. After incubation, HCAEC were washed three times with Tyrode solution containing 3.6% bovine serum albumin to remove unincorporated [3H]arachidonic acid. EC were incubated at 37°C for 15 min before implementation of the experimental conditions. At the end of the stimulation period, the surrounding medium was removed to a scintillation vial and represented the amount of radiolabeled arachidonic acid released from the HCAEC during the stimulation interval. The amount of radiolabeled arachidonic acid remaining in the endothelial monolayer was measured by adding 1 ml of 10% SDS, removing the cells from the culture well by scraping, and adding them to a scintillation vial. Radioactivity in both the surrounding medium and EC was quantified by liquid scintillation spectrometry.

Measurement of PAF production. HCAEC grown in 34-mm culture dishes (Corning) were washed twice with Hanks' balanced salt solution containing (in mM) 135 NaCl, 0.8 MgSO4, 10 HEPES (pH 7.4), 1.2 CaCl2, 5.4 KCl, 0.4 KH2PO4, 0.3 Na2HPO4, and 6.6 glucose. Cells were incubated with 10 µCi [3H]acetic acid/well for 20 min. After stimulation with tryptase, lipids were extracted from the cells using the method of Bligh and Dyer (1). The chloroform layer was concentrated by evaporation under N2, resuspended in 9:1 CHCl3/CH3OH, applied to a silica gel 60 TLC plate, and developed in chloroform-methanol-acetic acid-water (50:25:8:4 vol/vol/vol/vol). The region corresponding to [3H]PAF was scraped, and radioactivity was quantified by liquid scintillation spectrometry. Loss of PAF during extraction and chromatography was corrected by adding a known amount of [14C]PAF as an internal standard.

P-selectin surface expression assay. HCAEC, grown to confluence in 16-mm culture dishes (Corning), were incubated with tryptase with or without BEL pretreatment in Hanks' buffer at 37°C. At the end of incubations, buffer was quickly removed and cells were immediately fixed with 1% paraformaldehyde and incubated overnight at 4°C. Cells were then washed three times with PBS and blocked with Tris-buffered saline containing 0.1% (vol/vol) Tween 20 supplemented with 0.8% (wt/vol) BSA and 0.5% (wt/vol) fish gelatin for 1 h at 24°C. Cells were incubated with goat anti-P-selectin antibody (1:50) for 60 min at 37°C, washed with blocking buffer, and incubated with horseradish peroxidase-conjugated rabbit anti-goat antibody (1:5,000) for 30 min at 24°C. After being washed, each well was incubated in the dark for 30 min with the 3,3',5,5'-tetramethylbenzidine liquid substrate system. Reactions were stopped by the addition of sulfuric acid, and color development was measured with a microtiter plate spectrophotometer at 450 nm.

Changes in HCAEC monolayer electrical resistance. HCAEC were grown on presterilized Transwell polycarbonate filters mounted in a chamber insert. Once the cells reached confluence as determined visually, resistance across the HCAEC monolayer was measured with an epithelial voltohmmeter (EVOM; World Precision Instruments, Sarasota, FL) and miniature Ag-AgCl electrodes. Readings in ohms were taken directly from the EVOM. The resistance of the filter with no cells was subtracted from the values for filters containing cells to assess the resistance of the monolayer itself.

Isolation of neutrophils from human umbilical cord blood. Units of human umbilical cord blood were obtained from the St. Louis Cord Blood Bank. Units were pooled, and 25 ml of the pooled product was layered over 25 ml of Polymorphprep (Axis-Shield PoC, Oslo, Norway) in a 50-ml conical tube. Tubes were spun at 500 g for 30 min at 20°C with no brake. The top band at the sample-medium interface consisting of mononuclear cells and the lower band of polymorphonuclear cells were removed and placed into a clean 50-ml conical tube. An equal volume of 0.5 N Hanks' solution was added to the cells in the 50-ml tube. Normal Hanks' solution was added to bring the total volume to 50 ml. Tubes were spun at 400 g for 10 min at 4°C. Supernatant was discarded, and the cell pellet was resuspended with 3 ml of 0.2% NaCl and incubated for 3 min at room temperature; 3 ml of cold 1.6% NaCl was added, and the solution was transferred to a 15-ml conical tube. Ice-cold normal Hanks' solution was added to bring the total volume to 15 ml. Cells were centrifuged at 175 g for 10 min at 4°C. Supernatant was removed, and cells were resuspended in 5 ml of ice-cold Hanks’ solution. An aliquot was taken for a cell count with a hemacytometer. Cells were centrifuged at 175 g for 10 min, and the supernatant was discarded. Neutrophils were resuspended in MEM + 10% FCS at 1 x 106 cells/ml.

Neutrophil adherence assay. HCAEC were grown to confluence on a 12-mm plate. Cells were washed twice with MEM + 10% FCS and 0.5 ml of neutrophil suspension (5 x 105 cells) in MEM + 10% FCS, and tryptase was added to each of the wells and incubated for 10 min at room temperature. Media and unbound neutrophils were removed and discarded. Plates were washed twice with prewarmed Dulbecco's phosphate-buffered saline, and 1 ml of 0.2% Triton X-100 was added to each well to lyse adherent neutrophils and HCAEC. Cell lysates were scraped from the plate and transferred to an Eppendorf tube. A 0.5-ml aliquot of neutrophil suspension was added to 0.5 ml of 0.2% Triton X-100 and used as the theoretical maximal binding sample, and 0.5 ml of deionized H2O and 0.5 ml of 0.2% Triton X-100 were used as the reference blank. Samples, theoretical maximal binding sample, and blank were sonicated (550 Sonic Dismembrator; Fisher Scientific, Pittsburgh, PA) for 10 s. To measure neutrophil peroxidase activity, 400 µl of cell lysate was transferred to a glass tube, and 1 ml of PBS, 1,200 µl of Hanks' buffer + BSA, 200 µl of 3,3'-dimethoxybenzidine, and 200 µl of 0.05% H2O2 were added. The cell lysate reaction mixture was incubated for 15 min at room temperature, and 200 µl of 1% NaN3 was added to stop the reaction. The absorbance was then measured with a 4050 UV/visible spectrophotometer (Biochrom, Cambridge, UK) at 460 nm.


    RESULTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
As shown in Fig. 1, we performed immunoblot analysis using primary antibodies specific for PAR-2 to demonstrate the presence of this receptor in HCAEC. Stimulation of HCAEC with increasing concentrations of tryptase resulted in a concentration-dependent increase in PLA2 activity measured with (16:0, [3H]18:1) plasmenylcholine in the absence of Ca2+ (4 mM EGTA); thus the activity represents iPLA2. Increased iPLA2 activity was observed with concentrations of tryptase >0.5 ng/ml, with an ED50 for the concentration curve of 8 ng/ml tryptase. Consistent increases in iPLA2 activity were observed at 20 ng/ml (7.8 ± 0.2 vs. 2.2 ± 0.2 nmol·mg protein–1·min–1; P < 0.01, n = 3); therefore, this concentration of tryptase was used in further experiments.



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Fig. 1. Immunoblot analysis of protease-activated receptor (PAR)-2 in the isolated membrane fraction from human coronary artery endothelial cells (HCAEC). The 54-kDa band demonstrates the presence of PAR-2.

 
To examine the effects of tryptase stimulation on iPLA2 activity, HCAEC were grown to confluence and incubated with tryptase, and iPLA2 activity was measured. As shown in Fig. 2, tryptase stimulation induced a time-dependent increase in iPLA2 activity that was detectable at 1 min, reaching a peak at 5–10 min and then declining to basal levels after 30 min. The maximal, approximately threefold increase in tryptase-stimulated iPLA2 activity was similar to that observed previously in thrombin-stimulated HCAEC (20). Tryptase-stimulated PLA2 activity in HCAEC is mediated by an iPLA2 isoform, as evidenced by the fact that enzyme activity is maximal in the absence of Ca2+ and is completely inhibited by the iPLA2-selective inhibitor BEL (Fig. 2). To demonstrate that tryptase-induced cleavage of PAR-2 is responsible for the increase in iPLA2 activity, HCAEC were treated with SLIGKV, representing the tethered ligand sequence for the human PAR-2 once the receptor is cleaved by tryptase (Fig. 2). SLIGKV induced a time-dependent increase in iPLA2 activity similar to that seen with tryptase stimulation. These results indicate that the tryptase-stimulated increase in iPLA2 activity is working through activation of PAR-2.



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Fig. 2. Calcium-independent PLA2 (iPLA2) activation after tryptase stimulation. {blacksquare}, iPLA2 activity in HCAEC on stimulation with 20 ng/ml tryptase for increasing time; {square}, iPLA2 activity in HCAEC treated with 100 µM SLIGKV;, tryptase-stimulated cells pretreated with bromoenol lactone (BEL; 5 µm, 10 min). Data represent means ± SE for 4 separate experiments. *P < 0.05, **P < 0.01 compared with control.

 
To determine whether tryptase-stimulated iPLA2 activity catalyzes endogenous membrane phospholipid hydrolysis, we measured arachidonic acid release and PAF production in tryptase-stimulated HCAEC in the absence and presence of BEL. As shown in Fig. 3, stimulation of [3H]arachidonic acid-labeled HCAEC with tryptase induced a significant time-dependent increase in [3H]arachidonic acid release (Fig. 3). Pretreatment with BEL (Fig. 3) resulted in complete inhibition of tryptase-stimulated arachidonic acid release, demonstrating that arachidonic acid release is mediated by iPLA2.



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Fig. 3. Arachidonic acid release after tryptase stimulation. {blacksquare}, HCAEC stimulated with 20 ng/ml tryptase; {square}, tryptase-stimulated cells pretreated with BEL (5 µM, 10 min); {blacktriangleup}, arachidonic acid release by unstimulated cells. Data represent means ± SE for 6 separate experiments. *P < 0.05, **P < 0.01 compared with control.

 
As shown in Fig. 4, confluent HCAEC monolayers were incubated with tryptase (peak of iPLA2 activity) and assayed for PAF production with or without BEL pretreatment. Cells not pretreated with BEL (Fig. 4) demonstrate a significant increase in PAF production after tryptase stimulation. Pretreatment with BEL (Fig. 4) completely inhibits the tryptase-stimulated production of PAF from HCAEC, demonstrating that tryptase-induced PAF production is mediated via iPLA2.



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Fig. 4. Platelet-activating factor (PAF) production in stimulated HCAEC. HCAEC monolayers were incubated with 20 ng/ml tryptase for 10 min (filled bars). BEL pretreatment (open bars; 5 µM, 10 min) inhibited tryptase-stimulated PAF production. Additionally, HCAEC monolayers were incubated with 100 µM SLIGKV for 10 min. Data represent means + SE for at least 6 separate experiments. **P < 0.01 compared with control.

 
Confluent monolayers of HCAEC were incubated with SLIGKV and assayed for PAF production. SLIGKV treatment resulted in an increase in PAF production, similar to that seen with tryptase (Fig. 4), indicating that tryptase stimulation of HCAEC leading to PAF production occurs through cleavage of PAR-2.

Together these results demonstrate that iPLA2 is activated by tryptase stimulation (Fig. 2) resulting in the release of arachidonic acid (Fig. 3) and PAF production (Fig. 4). Tryptase stimulation of HCAEC leading to these biochemical responses occurs via cleavage of PAR-2 on the EC surface (Figs. 2 and 4). Additionally, pretreatment with BEL to inhibit iPLA2 activity (Fig. 2) results in the inhibition of both arachidonic acid release (Fig. 3) and PAF production (Fig. 4), demonstrating that tryptase-induced arachidonic acid release and PAF production are mediated via iPLA2-catalyzed hydrolysis of endogenous membrane phospholipids.

P-selectin expression on the EC surface was shown previously to be upregulated by PAR-1 activation; however, the combined role of PAR-2 and iPLA2 activation in P-selectin expression has not been previously examined. Therefore, we wished to determine whether tryptase treatment of the HCAEC monolayer also induced cell surface expression of P-selectin by a pathway dependent on iPLA2 activation. As shown in Fig. 5, cells were stimulated with tryptase with or without BEL pretreatment. Tryptase stimulation alone induced an increase in P-selectin expression. Pretreatment with BEL (Fig. 5) completely inhibited the increase in P-selectin cell surface expression in tryptase-stimulated cells. These results show that tryptase-activated HCAEC exhibit increased cell surface expression of P-selectin, occurring at least in part because of an iPLA2-dependent mechanism, potentially enhancing the binding of inflammatory cells bearing P-selectin glycoprotein ligand (PSGL)-1 to the HCAEC layer.



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Fig. 5. Cell surface P-selectin expression after tryptase stimulation. Filled bars, cell surface expression of P-selectin on HCAEC incubated with 20 ng/ml tryptase for 10 min; open bars, BEL pretreatment (5 µM, 10 min). Data represent means + SE for at least 6 separate cell cultures. abs, Absorbance. **P < 0.01 compared with control.

 
In contrast to the reversible interaction between EC P-selectin and PMN PSGL-1, EC PAF engages cognate PAF receptors on inflammatory cells, leading to inside-out activation of PMN integrins and high-affinity PMN binding to the endothelium. Because tryptase stimulation of HCAEC leads to increased PAF production and cell surface expression of P-selectin, we determined whether these events promote tight adhesion of circulating neutrophils to the endothelium mediated by PMN integrins activated by HCAEC PAF. Neutrophils were isolated from human umbilical cord blood and incubated with a confluent monolayer of HCAEC stimulated with tryptase (see METHODS). A peroxidase assay was performed on the cell lysate to determine the percentage of adherent neutrophils. As shown in Fig. 6, tryptase stimulation of HCAEC monolayers increased neutrophil adherence 2.4-fold over levels of neutrophil adhesion to unstimulated HCAEC. Pretreatment of the HCAEC with BEL significantly inhibited neutrophil adherence after tryptase stimulation, demonstrating the importance of iPLA2-dependent regulation of PMN adherence.



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Fig. 6. Effect of tryptase stimulation on the adhesion of neutrophils to HCAEC. These data demonstrate a 2.5-fold increase in polymorphonuclear neutrophil adhesion on stimulation of HCAEC with 20 ng/ml tryptase alone for 10 min (filled bars). Open bars, pretreatment with BEL (5 µM, 10 min). Results represent means + SE for at least 4 separate cell cultures. **P < 0.01 compared with control.

 
In addition to enhanced inflammatory cell binding, confluent PAR-activated EC may detach from neighboring cells, presumably in preparation for inflammatory cell transendothelial migration (6). To determine the effect of tryptase on EC monolayer barrier properties, HCAEC were grown to confluence on a Transwell insert and then treated with increasing concentrations of tryptase, and the resultant changes in electrical resistance across the EC monolayer were measured (Fig. 7). Tryptase stimulation resulted in a time-dependent decrease in electrical resistance across the HCAEC monolayer that started at ~30 min. After ~2 h, transendothelial resistance started to return toward unstimulated values.



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Fig. 7. Electrical resistance of HCAEC on stimulation with 2, 20, or 200 ng/ml tryptase. These data demonstrate the time-dependent decrease in electrical resistance on tryptase stimulation. Results represent means ± SE for at least 8 separate cell cultures.

 

    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In a recent study (20), we demonstrated that thrombin stimulation of HCAEC PAR-1 activates a novel membrane-associated iPLA2 that selectively hydrolyzes plasmalogen phospholipids. The accelerated plasmalogen phospholipid hydrolysis results in the production of several biologically active phospholipid metabolites, including arachidonic acid, prostacyclin, lysoplasmenylcholine, and PAF, that may contribute to inflammation. These responses were all inhibited by pretreatment with the iPLA2-selective inhibitor BEL. Additionally, previously published studies from our laboratory (19) demonstrate that thrombin stimulation of PAR-1 in HCAEC is specifically coupled to activation of iPLA2 and that arachidonic acid production and PAF synthesis in HCAEC proceed exclusively through a pathway in which iPLA2 represents the initial, rate-controlling step.

In this study, we have demonstrated that incubation of HCAEC with tryptase reproduces a similar pattern of iPLA2 activation, arachidonic acid release, and PAF production, suggesting that in HCAEC activation of PAR-2 may couple to similar intracellular pathways associated with inflammation. These data agree with a previously published study that suggests that the presence of multiple PARs on the EC surface serves to extend the number of proteases to which the cells respond rather then being coupled to different intracellular responses (22). These data also suggest that PAR may play important roles in both the acute and chronic inflammatory behavior of both endothelial and epithelial cells that form the defensive barriers of the body (26).

PAF is a membrane phospholipid-derived inflammatory mediator that plays a role in propagating the inflammatory response, and it promotes aggregation, chemotaxis, granule secretion, and oxygen radical generation from PMN, and the adherence of PMN to the endothelium. PAF also increases the permeability of the HCAEC monolayer and stimulates smooth muscle contraction. Previous studies demonstrate that thrombin stimulation of HCAEC activating PAR-1 leads to an increase in iPLA2 activity (19). Accordingly, PAR activation of iPLA2 coupled with PAF production may be a key event in inflammatory cell recruitment, control of thrombosis, fibrinolysis, and vascular permeability and vascular remodeling by mast cell-derived effectors such as tryptase. Increases in HCAEC PAF expression and alterations in the phospholipid composition of the HCAEC membrane enhance inflammatory cell adherence and transmigration, respectively, allowing these cells access to sites of vascular injury (21). Additionally, increased production of eicosanoids synthesized from arachidonic acid is involved in regulating the body's response to inflammation (7, 25).

P-selectin is an adhesion molecule involved in the early stages of tethering of freely circulating cells to an activated endothelium. The presence of P-selectin on an activated EC layer plays an essential role in the initiation of a tentative adhesive interaction between the circulating inflammatory cell and activated EC layer (21). Studies by Torres et al. (28) have demonstrated the ability of mast cell mediators to increase P-selectin expression in canine carotid artery EC. Although their findings demonstrate that P-selectin expression occurs via a histamine-independent mechanism, they do not attribute this effect to a specific mast cell mediator, allowing for the possibility that tryptase is integral to this process.

The enhanced presence of HCAEC-associated PAF has been shown to cause transient adherence of neutrophils, which bear the PAF receptor, to the HCAEC (10, 11, 18, 23). Prescott et al. (27) correlated the adhesion of neutrophils to thrombin-activated endothelium with PAF synthesis and expression on the surface of EC. In these experiments, we have demonstrated a 2.4-fold increase in PMN adhesion on stimulation of HCAEC with tryptase (Fig. 6). Increased PAF production and expression of P-selectin on the surface of the HCAEC monolayer may similarly lead to the recruitment of other inflammatory cells bearing the PAF receptor and/or PSGL-1, allowing for their binding to PAF and P-selectin expressed on the surface of the EC. In these experiments, we have shown that tryptase stimulation results in a time-dependent decrease in electrical resistance across the HCAEC monolayer (Fig. 7) that returns to basal levels after ~2 h. The decrease in resistance suggests that tryptase stimulation increases HCAEC monolayer permeability, possibly through a decrease in the number of stable tight junctions, whereas the return to normal levels of electrical resistance may indicate the reformation of stable tight junctions. Doukas et al. (6) demonstrated that histamine treatment of EC, which increases the permeability of the EC, increases the number of PMN migrating across the endothelium. We hypothesize that decreased resistance corresponds to an increase in the permeability between adjacent EC in the monolayer and may facilitate the passage of inflammatory cells across the EC monolayer after their initial adherence to the EC surface.

The discovery of mast cells, activated T lymphocytes, and macrophages in atherosclerotic lesions, the detection of human PMN antigen class II expression, and the finding of local secretion of several cytokines all suggest the involvement of immune and inflammatory mechanisms in the pathogenesis of atherosclerosis (12, 17, 29). The recent identification of increased mast cell numbers in atherosclerotic plaques suggests these cells may play an important role in the evolution of atherosclerosis (3, 1416, 24). As reviewed by Galli et al. (9), mast cells are known to be activated during innate immune responses, leading to the release of their mediators, such as tryptase, which can, as demonstrated in the studies presented here, activate iPLA2, resulting in the production of the inflammatory mediators arachidonic acid and PAF. Our data suggest that these biochemical events mediate the attachment and endothelial transmigration of circulating inflammatory cells and may represent a mechanism whereby these cells may accumulate in the vascular bed. We propose that activation of the endothelium via mast cell tryptase, leading to the accumulation of inflammatory cells in the area of vascular injury, may be one of the early inflammatory events involved in the progression to plaque formation. The unique ability of the mast cell to produce and release tryptase may provide a mechanism by which these cells propagate the inflammatory response.

In conclusion, we have shown that tryptase activates HCAEC iPLA2, leading to increases in PAF production, cell surface P-selectin expression, and recruitment of neutrophils to the HCAEC monolayer. The presence of PAF and P-selectin on the HCAEC surface provides a mechanism for any circulating cell expressing the P-selectin ligand and PAF receptor to adhere to the activated HCAEC monolayer. We hypothesize that tryptase-producing mast cells found in areas of atherosclerotic injury become activated, leading to the release of tryptase and the subsequent recruitment of inflammatory cells and thus propagating the inflammatory response.


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 ABSTRACT
 METHODS
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 GRANTS
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This work was supported in part by National Heart, Lung, and Blood Institute Grant HL-68588 (J. McHowat) and the American Heart Association, Heartland Affiliate (J. McHowat).


    ACKNOWLEDGMENTS
 
The authors acknowledge the St. Louis Cord Blood Bank for the supply of human umbilical cord blood units for neutrophil isolation.


    FOOTNOTES
 

Address for reprint requests and other correspondence: J. McHowat, Dept. of Pathology, Saint Louis Univ. School of Medicine, 1402 S. Grand Blvd., St. Louis, MO 63104 (e-mail: jane.mchowat{at}tenethealth.com)

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.


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