Differential Ca2+ signaling by thrombin and protease-activated receptor-1-activating peptide in human brain microvascular endothelial cells

Yuri V. Kim,1 Francescopaolo Di Cello,1 Coryse S. Hillaire,2 and Kwang Sik Kim1

1Division of Pediatric Infectious Diseases and 2Department of Neurology, Johns Hopkins University, School of Medicine, Baltimore, Maryland 21287

Submitted 22 April 2003 ; accepted in final form 21 August 2003


    ABSTRACT
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Thrombin and related protease-activated receptors 1, 2, 3, and 4 (PAR1–4) play a multifunctional role in many types of cells including endothelial cells. Here, using RT-PCR and immunofluorescence staining, we showed for the first time that PAR1–4 are expressed on primary human brain microvascular endothelial cells (HBMEC). Digital fluorescence microscopy and fura 2 were used to monitor intracellular Ca2+ concentration ([Ca2+]i) changes in response to thrombin and PAR1-activating peptide (PAR1-AP) SFFLRN. Both thrombin and PAR1-AP induced a dose-dependent [Ca2+]i rise that was inhibited by pretreatment of HBMEC with the phospholipase C inhibitor U-73122 and the sarco(endo)plasmic reticulum Ca2+-ATPase inhibitor thapsigargin. Thrombin induced transient [Ca2+]i increase, whereas PAR1-AP exhibited sustained [Ca2+]i rise. The PAR1-AP-induced sustained [Ca2+]i rise was significantly reduced in the absence of extracellular calcium or in the presence of an inhibitor of store-operated calcium channels, SKF-96365. Restoration of extracellular Ca2+ to the cells that were initially activated by PAR1-AP in the absence of extracellular Ca2+ resulted in significant [Ca2+]i rise; however, this effect was not observed after thrombin stimulation. Pretreatment of the cells with a low thrombin concentration (0.1 nM) prevented [Ca2+]i rise in response to high thrombin concentration (10 nM), but pretreatment with PAR1-AP did not prevent subsequent [Ca2+]i rise to high PAR1-AP concentration. Additionally, treatment with thrombin decreased transendothelial electrical resistance in HBMEC, whereas PAR1-AP was without significant effect. These findings suggest that, in contrast to thrombin, stimulation of PAR1 by untethered peptide SFFLRN results in stimulation of store-operated Ca2+ influx without significantly affecting brain endothelial barrier functions.

store-operated calcium influx; desensitization; transendothelial electrical resistance; digital imaging


BRAIN MICROVASCULAR ENDOTHELIAL cells represent a key component of the blood-brain barrier (BBB). Compared with endothelial cells of nonbrain origin few molecules can cross the BBB and any increases in permeability of the barrier are usually considered detrimental. Many pathological conditions like acute inflammation, atherosclerosis, and stroke are associated with an increased generation of thrombin (9, 11, 17, 26, 28, 51). The endothelial cells represent one of the main targets of thrombin action, and thrombin has been shown to exhibit a variety of effects on endothelial cells, e.g., production of cytokines and chemokines, vasodilation, and expression of adhesion molecules (9, 11, 28). Thrombin-mediated effects on endothelial cells involve activation of a unique group of protease-activated receptors (PARs) (10, 21, 28). PARs belong to the G protein-coupled receptors, and currently four members of the group are known: PAR1, PAR2, PAR3, and PAR4. Thrombin activates PAR1, PAR3, and PAR4, whereas PAR2 is activated by other proteases, trypsin and tryptase (10, 21, 28). Thrombin activation of the best-studied PAR1 is associated with cleavage of the extracellular NH2-terminal domain of the receptor, and subsequent exposure of a new NH2 terminus, so-called tethered ligand, can interact with one of the extracellular loops of PAR1 and thus induce signal transduction (10). Subsequent studies showed that PAR1 could be activated not only by tethered ligand but also with synthetic peptides, analogs of the last 5–14 amino acids of the new NH2 terminus, such as SFFLRN (3, 5, 10, 19, 21). The same mechanisms of activation apply to PAR2, -3, and -4 (10, 21, 28). However, several recent studies demonstrated that activation of PARs by tethered ligands could differ from the effects induced by the synthetic, untethered analog peptides (1, 5, 12, 20, 22). These data raise the interesting possibility of differential activation of PAR1 by different agonists. PARs are widely expressed in the brain (42); however, the presence and mechanisms of activation of PARs in brain microvascular endothelial cells are poorly understood. For example, the presence of PAR1, PAR2, and PAR3 mRNA and the intracellular Ca2+ concentration ([Ca2+]i) signal induced by thrombin and PAR1-activating peptide (PAR1-AP) have been shown in primary cultures of rat brain microvascular cells (4). In another report, thrombin induced [Ca2+]i rise and extracellular Ca2+ influx in human cerebral capillary endothelial cells (27).

In the present study we examined whether PARs are expressed in primary cultures of human brain microvascular endothelial cells (HBMEC) that constitute the BBB and performed comparative analysis of the mechanisms of thrombin- and PAR1-AP-induced Ca2+ signaling and endothelial barrier dysfunction. We showed the presence of mRNA and surface expression of all four PARs in HBMEC. We also found that, in contrast to thrombin, the activation of PAR1 by the untethered agonist peptide SFFLRN resulted in stimulation of massive extracellular Ca2+ influx through store-operated calcium channels (SOCCs). However, the peptide failed to affect transendothelial electrical resistance (TER) across the HBMEC monolayer, whereas thrombin caused a decrease in TER.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Materials. Thrombin, thapsigargin, and HEPES were purchased from Sigma (St. Louis, MO). U-73122 and SKF-96365 were from Calbiochem (San Diego, CA). M199, fetal bovine serum (FBS), and Hanks' balanced salt solution (HBSS) were from GIBCO (Grand Island, NY). PAR1, PAR2, PAR3, and PAR4 peptides (SFFLRN, SLIGKV, TFRGAP, and GYPGKF, respectively) were from Bachem (Bachem USA, Torrance, CA). Another thrombin receptor-activating peptide (TRAP; Ala-pFluoro-Phe-Arg-Cha-HomoArg-Tyr-NH2) was purchased from Neosystem (Strasbourg, France). Rat tail collagen was purchased from BD Bioscience (Bedford, MA). Acetylated low-density lipoprotein (LDL) labeled with 1,1'-dioctadecyl-3,3,3',3'-tetramethylindo-carbocyanine perchlorate was obtained from Biomedical Technologies (Stoughton, MA). Fura 2-AM and Pluronic 123 were from Molecular Probes (Eugene, OR).

Cell culture. Primary HBMEC were isolated and characterized as described previously (43, 44). The quality of the endothelial cells was assessed by evaluation of specific labeling of the cells with fluorescent acetylated LDL (DiI-Ac-LDL). More than 95% of the cells showed fluorescent staining with the label. On cultivation on collagen-coated Transwell inserts these HBMEC exhibited TER of 300–600 {Omega}·cm2 (34, 43), a unique property of the brain microvascular endothelial monolayer compared with systemic vascular endothelium. The frozen stock of HBMEC between passages 8 and 13 was thawed and cultured in M199 supplemented with 10% (vol/vol) heat-inactivated FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin in 25-cm2 flasks. The cell cultures were maintained at 37°C in a humidified atmosphere of 5% CO2-95% air. After reaching confluence, the cells were detached with 0.05% trypsin-0.5 mM EDTA solution, resuspended in fresh M199 medium, and seeded on glass coverslips (Fisher Scientific, Houston, TX) coated with 0.1% rat tail collagen. Twelve to twenty hours before experiments, coverslips with cells were placed in lowserum M199 medium with 0.5–1% FBS.

Reverse transcriptase-polymerase chain reaction. Total RNA was extracted from HBMEC cultures grown on collagen-coated 60-mm dishes with a RNeasy Mini Kit (Qiagen, Valencia, CA) by applying the on-column DNase treatment according to the manufacturer's instructions. The amount and quality of the RNA were verified by measuring the absorbance at 260 and 280 nm. Oligo(dT)-primed reverse transcription of RNA was performed with the SuperScript First-Strand Synthesis System for reverse transcriptase-polymerase chain reaction (RT-PCR) (Invitrogen, Carlsbad, CA), using 1 or 0.1 µg of RNA for each reaction.

PCR amplifications were performed from 2 µl of each cDNA sample with QuantumRNA {beta}-actin primers (Ambion, Austin, TX) or specific primers from Asokananthan et al. (2). The following forward and reverse primers were used for amplifying human PARs: PAR1: forward 5-TGTGAACTGATCATGTTTATG-3', reverse 5'-TTCGTAAGATAAGAGATATGT-3' (PCR product, 708 bp); PAR2: forward 5'-AGAAGCCTTATTGGTAAGGTT-3', reverse 5'-AACATCATGACAGGTCGTGAT-3' (PCR product, 582 bp); PAR3: forward 5'-CTGATACCTGCCATCTACCTCC-3', reverse 5'-AGAAAACTGTTGCCCACACC-3' (PCR product, 382 bp); PAR4: forward 5'-ATTACTCGGACCCGAGCC-3', reverse 5'-TGTAAGGCCCACCCTTCTC-3' (PCR product, 392 bp). The PCR program consisted of one preincubation at 94°C for 2 min and 40 cycles at 94°C for 30 s, 55°C for 30 s (50°C for 1 min for PAR1), and 68°C for 1 min (3 min for PAR1). All PCR reactions were performed with a Robocycler Gradient 40 with a heated lid (Stratagene, La Jolla, CA) in 50 µl of 1x PCR buffer, 1.5 mM MgCl2, each primer at 0.2 µM, 200 µM dNTP, and 1 U of Taq DNA polymerase (Invitrogen). Amplification mixtures were analyzed by agarose gel electrophoresis.

Immunofluorescent staining of PARs. Primary HBMEC grown on coverslips were fixed in 4% paraformaldehyde for 10 min at room temperature. The cells were washed three times with PBS, 5 min each, then blocked for 1 h in a PBS solution containing 4% donkey serum (Sigma) and 0.3% Triton X-100. This was followed by 4°C overnight incubation with 1:200 dilution of the primary PAR antibody (goat anti-human PAR1, RDI; mouse anti-human PAR2 and PAR4 and rabbit anti-human PAR3, Santa Cruz Biotechnology). Normal rabbit IgG, mouse IgG, and goat IgG were used as negative controls, and they did not show any fluorescence increase above background. Cells were washed three times in PBS, 5 min each, and incubated for 2 h at room temperature with secondary antibodies diluted 1:500 in PBS (Alexa 488-conjugated anti-goat or Cy3-conjugated anti-rabbit and Cy3-conjugated anti-mouse or Alexa 488-conjugated anti-mouse; Jackson Lab). Coverslips with the cells were again washed three times in PBS before being mounted onto the stage of a fluorescent microscope.

[Ca2+]i measurements. HBMEC were grown in M199 supplemented with 10% FBS on 22-mm collagen-coated square glass coverslips until at least 80% confluence. The coverslips were washed with HEPES-buffered HBSS containing (mM) 137 NaCl, 4.2 NaHCO3, 0.4 Na2HPO4, 5.4 KCl, 0.4 KH2PO4, 1. 3 CaCl2, 0.5 MgCl2, 0.4 MgSO4, 5.6 D-glucose, 2 Na-pyruvate, and 15 HEPES buffered at pH 7.4 and incubated for 40 min with 3 µM fura 2-AM and 0.04% Pluronic 123 in the dark at room temperature. After loading, the cells were washed from extracellular fura 2-AM and incubated in the same medium for an additional 20 min. Cells loaded with fura 2 were mounted in the recording chamber (Warner Instruments, Hamden, CT) on the microscope stage, and fluorescence images were captured with an Olympus fluorescence microscopy system (Olympus America, Melville, NY) equipped with an inverted Olympus microscope IX-70, a cooled charge-coupled device camera OlymPix (model TE3/A/S; AstroCam), a x40, 1.3-numerical aperture oil-immersion objective, and a computer-controlled Sutter filter wheel (Sutter Instrument, Novato, CA). Before fluorescence measurements started, 25–40 regions of interest representing individual cells were selected on the field of view to control the experiments. Fura 2 fluorescent images were captured at 2-s intervals by alternating excitation of cells at 340 and 380 nm wavelengths and reflection off dichroic mirror with a cutoff wavelength at 510 nm and band-pass emission filtering centered at 530 nm. The real-time fluorescent images were displayed on a monitor and stored on a hard drive for subsequent detailed analysis with UltraView software (Olympus America). Changes in [Ca2+]i were expressed as the 340- to 380-nm ratio (18). Ca2+-free solutions were made with HEPES-buffered Ca2+-free HBSS with an additional 0.1 mM EGTA.

Transendothelial electrical resistance. To study endothelial barrier dysfunction, a new special ECIS (electric cell-substrate impedance sensing) system (Applied Biophysics, Troy, NY) for measuring characteristics of cell growth, attachment/spreading, and barrier function of confluent cell layers (45) was used to measure the changes in TER during exposure to thrombin and PAR1-AP. HBMEC were directly seeded on a collagen-coated eight-well gold electrode array in M199 supplemented with 10% FBS. Each well had one active electrode (250-µm diameter) and a large counterelectrode. Both electrodes were connected to a phase-sensitive lock-in amplifier. The electrodes were fed with a constant current of 1 µA supplied by a 1-V, 4,000-Hz AC signal. Each well contained 400 µl of the medium. Initial resistance of electrodes in M199 medium was ~2,000 {Omega}. After reaching maximal, steady-state readings of transendothelial resistance (~15,000 {Omega}) that meant maximal confluence, cells were additionally incubated overnight in low-serum (1% FBS) M199 medium. Before the experiment, HBMEC monolayers were incubated for 2 h in HEPES-buffered HBSS and then thrombin- and PAR1-AP-induced changes in resistance of endothelial monolayers were monitored. Data on traces are presented as changes in resistance normalized to time zero before additions of thrombin and PAR1-AP.

Statistical analysis. Experimental data are presented as means ± SE of 4–10 independent experiments. Probability values of P < 0.05 according to unpaired Student's t-test were considered significant.


    RESULTS
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Expression of PARs in HBMEC. Previous studies using RT-PCR showed that primary cultures of rat brain capillary endothelial cells express PAR1, PAR2, and PAR3, but the presence of PAR4 was undetermined (4). It is unknown whether all PARs are expressed in HBMEC. To assess the presence of PARs in HBMEC, we used qualitative RT-PCR with specific primers (2) to amplify PAR1, PAR2, PAR3, and PAR4 mRNA from the total RNA isolated from HBMEC grown in M199 medium on collagen-coated culture dishes. We showed that mRNAs of all four PARs were detected and the sizes of the PCR products were consistent with the expected sizes based on primers (Fig. 1A). Immunofluorescent staining of nonpermeabilized HBMEC also showed surface expression of all four PARs (Fig. 1, B–E).



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Fig. 1. Expression of protease-activated receptors PAR1, PAR2, PAR3, and PAR4 in primary cultures of human brain microvascular endothelial cells (HBMEC). A: reverse transcriptase-polymerase chain reaction (RT-PCR) of PARs. Total mRNA were isolated from HBMEC grown on collagencoated culture dish in M199 containing 10% fetal bovine serum (FBS). RT-PCR amplification of PAR-specific primers was performed with the SuperScript First-Strand Synthesis System for RT-PCR. First and last columns: GeneRuler 100-bp Ladder Plus. +, RT-PCR amplification; –, minus RT control. B–E: surface expression of PAR1, -2, -3, and -4 on HBMEC. Immunofluorescent staining (green) of nonpermeabilized HBMEC with PAR1, -2, -3, and -4 antibodies is shown. Bar, 40 µm.

 

Effects of thrombin and PAR-AP on [Ca2+]i of HBMEC. We next examined and compared [Ca2+]i changes in HBMEC in response to thrombin and PAR-AP. Figure 2 shows typical changes of [Ca2+]i in HBMEC during exposure to various concentrations of thrombin (Fig. 2, A–C) and PAR1-AP SFLLRN (Fig. 2D). Thrombin-induced [Ca2+]i rise was rapid and transient, whereas PAR1-AP induced a rapid, biphasic, and prolonged [Ca2+]i rise. Dose-response curves showed that thrombin induced maximal [Ca2+]i rise at ~10 nM (or 1 NIH U/ml), whereas no [Ca2+] increase was observed at concentrations <100 pM (0.01 NIH U/ml) (Fig. 3A). In contrast, PAR1-AP induced a dose-dependent [Ca2+] rise up to 500 µM and no [Ca2+]i changes were observed at <1 µM (Fig. 3B). Similar differences in cell sensitivity to thrombin and PAR1-AP were observed in other types of cells, such as pulmonary endothelial cells, astrocytes, and smooth muscle cells (21, 28), and probably reflect differences in availability of PAR1 binding sites to thrombin-unmasked tethered and untethered PAR1-AP peptides. We tested [Ca2+] responses of HBMEC to activating peptides of all four PARs. As shown in Fig. 4, only PAR1-AP and PAR2-AP were able to induce [Ca2+]i changes, whereas PAR3-AP and PAR4-AP failed to induce any significant [Ca2+]i changes in HBMEC even at up to 500 µM concentrations. This data suggests that PAR3 and PAR4 receptors may not be directly connected to Ca2+ signaling pathways in HBMEC.



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Fig. 2. Thrombin- and PAR1-induced changes of intracellular Ca2+ concentration ([Ca2+]i) in HBMEC. Cells were grown on coverslips as described in EXPERIMENTAL PROCEDURES, loaded with fura 2-AM, and challenged with various concentrations of thrombin and PAR1-activating peptide (PAR1-AP) SFFLRN. A–C: where indicated by arrows, 1, 10, and 100 nM thrombin was added, respectively. D: where indicated by horizontal lines, 1, 10, and 100 µM PAR1-AP (P1) SFFLRN was added sequentially. Representative traces of single cells of 4–8 experiments are presented. In each experiment, 25–40 cells were selected to measure [Ca2+]i changes, expressed as the 340- to 380-nm ratio of fura 2 fluorescence.

 


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Fig. 3. Concentration-dependent effects of thrombin and PAR1-AP on [Ca2+]i in HBMEC. Average peak [Ca2+]i changes induced by thrombin (A) and PAR1-AP (B) of 20–40 cells in each experiment of 3–8 independent preparations were pooled. Data are expressed as means ± SE.

 


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Fig. 4. PAR-AP-induced cytosolic [Ca2+] responses in HBMEC. Cells were loaded with fura 2-AM and treated with PAR1-AP, PAR2-AP, PAR3-AP, and PAR4-AP (SFFLRN, SLIGKV, TFRGAP, and GYPGKF, respectively). Horizontal lines show additions: 50 µM PAR2-AP (P2) and 50 µM PAR1-AP (P1) (A); 200 µM PAR3-AP (P3), 10 µM and 500 µM PAR1-AP (P1) (B); and 200 µM PAR4-AP (P4) and 100 µM PAR1-AP (P1) (C). Unmarked arrow in C indicates addition of 10 µM ATP to show functional integrity of the cells. Representative traces of single cells of at least 3 independent experiments are presented. In each experiment, [Ca2+]i changes in >20 cells were measured.

 

Effects of inhibitors of phospholipase C and sarco(endo)-plasmic reticulum Ca2+-ATPase. PAR1 belongs to the seven-transmembrane spanning G protein-coupled receptor family, and its activation is associated with mobilization of intracellular Ca2+ (10, 28). To investigate the source of PAR1-induced [Ca2+]i increase in HBMEC, we tested the effects of inhibitors of phospholipase C (PLC) and sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA), U-73122 and thapsigargin, respectively (Fig. 5). Preincubation of HBMEC with 10 µM U-73122 for 10 min prevented subsequent PAR1-AP-induced and thrombin-induced (data not shown) [Ca2+]i rise (Fig. 5A). Figure 5B shows that after Ca2+ discharge from intracellular Ca2+ stores by 1 µM thapsigargin, subsequent addition of 100 µM PAR1-AP did not induce any additional increase of [Ca2+]i. Similarly, thrombin did not induce any [Ca2+]i changes in thapsigargin-treated cells in the absence of extracellular Ca2+ (Fig. 5C). These data suggest the critical importance of a PLC- and thapsigargin-sensitive Ca2+ pool during PAR1 activation with thrombin or PAR1-AP.



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Fig. 5. Pretreatment with thapsigargin and U-73122 prevented PAR1-AP-induced [Ca2+]i changes in HBMEC. A: cells were incubated with 10 µM U-73122 for 10 min and challenged with 500 µM PAR1-AP (P1; horizontal line) and where indicated by arrow with 1 µM thapsigargin (Thap). B and C: where indicated, cells were treated with 1 µM thapsigargin and then with 500 µM PAR1-AP (P1) in the presence (B) and absence (C) of extracellular Ca2+ (). Representative traces of 3 independent experiments are presented. Similar traces were observed when 100 nM thrombin was added instead of PAR1-AP.

 

Role of extracellular Ca2+ influx. As indicated above, in contrast to thrombin, PAR1-AP induced biphasic [Ca2+]i increase with a prolonged phase of an increased [Ca2+]i. This different response can be attributed to the possibility that PAR1-AP-induced activation of HBMEC may result in greater extracellular Ca2+ influx compared with thrombin. The area under the [Ca2+]i peak curve and/or time required to return to basal levels of [Ca2+]i can serve as an indirect measure of estimating magnitude of Ca2+ changes. The larger area (and/or the longer time) means more Ca2+ entering the cells and/or less Ca2+ efflux. For instance, half-peak widths of thrombin- and PAR1-AP induced [Ca2+]i were 44 ± 4 and 89 ± 12 s (n = 4; P < 0.01) for 10 nM thrombin and 500 µM PAR1-AP, respectively (Fig. 6A). The differences between thrombin and PAR1-AP became even more significant when full recovery time of [Ca2+]i was estimated: 106 ± 16 and 297 ± 41 s, respectively (n = 5; P < 0.005). Similar findings were obtained when the total areas of thrombin- and PAR1-AP-induced peak [Ca2+]i rise were compared. In normal HBSS medium these values were 81 ± 19 and 207 ± 24 relative area units (n = 7–10; P < 0.001), respectively, for 10 nM thrombin and 500 µM PAR1-AP (Fig. 6B). Considering the possibility that the effect of PAR1-AP could be due to activation of PAR2 (24), we pretreated HBMEC with trypsin or PAR2-AP SLIGKV and then compared Ca2+ responses to thrombin and PAR1-AP. Such pretreatment with trypsin or PAR2-AP did not eliminate the prolonged phase of PAR1-AP-induced [Ca2+]i increase compared with thrombin (data not shown; see also Fig. 4A). The kinetics of calcium responses did not significantly change when we used another artificial PAR1-AP, Ala-pFluoro-Phe-Arg-Cha-HomoArg-Tyr-NH2 (data not shown). These findings suggest that PAR1-AP-induced extracellular Ca2+ influx was not due to nonspecific activation of PAR2. Removal of extracellular Ca2+ significantly reduced peak area under the curve. For example, for PAR1-AP-induced peak [Ca2+]i increase this value became 148 ± 14 (n = 7), compared with 207 ± 14 obtained in the presence of extracellular Ca2+. Removal of extracellular Ca2+ also decreased the time and decreased the area of cytoplasmic [Ca2+]i recovery after thrombin stimulation (Fig. 6B). Additionally, thrombin induced much less Ca2+ loss from the thapsigargin-sensitive Ca2+ pool compared with PAR1-AP (Fig. 6, C and D). In the absence of extracellular Ca2+, the area of the thapsigargin-, thrombin-, and PAR1-AP-induced [Ca2+]i rise was 113 ± 9, 27 ± 4, and 70 ± 6, respectively. After thrombin and PAR1-AP stimulation the thapsigargin-sensitive Ca2+ pool was 93 ± 5 and 21 ± 3, respectively (means ± SE; n = 40–50 cells; P < 0.001; Fig. 6E). These data supported the hypothesis that activation with PAR1-AP results in an increased influx of extracellular Ca2+ compared with thrombin because of increased Ca2+ depletion from intracellular Ca2+ stores. The opening of calcium channels during PAR1-AP activation was apparent after readdition of extracellular Ca2+, which resulted in prolonged increase of [Ca2+]i (Fig. 7A). In contrast, after thrombin activation of HBMEC readdition of extracellular Ca2+ did not cause significant increase of [Ca2+]i (Fig. 7B). One of the possible mechanisms of Ca2+ entry is related to activation of SOCCs, induced by Ca2+ store depletion and a capacitative Ca2+ entry (CCE) mechanism (38, 48). To test this, we used a specific blocker of SOCCs, SKF-96365. As shown in Fig. 7C, preincubation of the cells with 50 µM SKF-96365 resulted in near elimination of PAR1-AP-induced sustained increase of [Ca2+]i. La3+, another known general inhibitor of calcium channels, also prevented the effect of readdition of extracellular Ca2+ on PAR1-AP-stimulated HB-MEC (Fig. 7D). The inhibitory effect of La3+ was observed at 10 µM, whereas complete inhibition of [Ca2+]i increase occurred at 1 mM. In contrast, verapamil, an inhibitor of voltage-activated calcium channels, was without any effect (data not shown).



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Fig. 6. PAR1-AP activation of HBMEC results in an increased Ca2+ discharge from thapsigargin-sensitive Ca2+ pool and a prolonged extracellular Ca2+ influx. Cells grown on coverslips were loaded with fura 2-AM and challenged with 10 and 500 µM PAR1-AP SFFLRN and 0.1 and 10 nM thrombin (Thr). A: half-time (T1/2) and total time (Ttotal) required for complete return of PAR1-AP- and thrombin-induced peak [Ca2+]i rise to prestimulatory level. Data are expressed as means ± SE; n = 4–9 independent experiments. *P < 0.05 and **P < 0.001 between 0.5 mM PAR1-AP and 10 nM thrombin. B: total area under PAR1-AP- and thrombin-induced peak [Ca2+]i rise. Cells were activated in the presence (+) and absence (–) of . Data are expressed as means ± SE; n = 3–8 experiments. In each experiment Ca2+ responses of >15 cells were averaged. *P < 0.05 between PAR1-AP-induced peak [Ca2+]i rise in the presence and absence of extracellular calcium; **P < 0.001 between 500 µM PAR1-AP- and 10 nM thrombin-induced [Ca2+]i increase. C and D: cells were treated where indicated with 10 nM thrombin (C) or 500 µM PAR1-AP (D) in the absence of extracellular Ca2+ and then 1 µM thapsigargin was added where indicated. E: total area of thapsigargin-induced [Ca2+]i rise (hatched bars) after thrombin and PAR1-AP stimulation in the absence of extracellular Ca2+. Data are means ± SE; n = 40–50 cells. **P < 0.001 between thrombin and PAR1-AP.

 


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Fig. 7. Activation of PAR1 receptor by PAR1-AP caused stimulation of extracellular Ca2+ influx through store-operated Ca2+ channels. Cells were challenged with 500 µM PAR1-AP (P1; A) or with 100 nM thrombin (Thr; B) in the absence of and then normal Hanks' balanced salt solution (HBSS) containing 1.2 mM Ca2+ was restored . C: cells were preincubated or sham-incubated with 50 µM SKF-96365, an inhibitor of store-operated calcium channels, and then challenged with 500 µM PAR1-AP. D: cells were treated with 500 µM PAR1-AP in the absence of and then incubated in normal HBSS containing Ca2+ and 1 mM La3+. Representative traces of single cells of 3 independent experiments are presented. In each experiment Ca2+ changes in >20 cells were measured. Arrows indicate addition of 10 µM ATP.

 

Desensitization of thrombin-induced [Ca2+]i changes. Previous reports showed that stimulation of endothelial cells of nonbrain origin with a lower dose of thrombin caused dramatic desensitization of PAR1 to subsequent stimulation with much higher concentrations of thrombin (24, 32). The mechanism of such desensitization remains unclear. In this series of experiments, we tested whether in HBMEC the same mechanism of desensitization may apply to thrombin and PAR1-AP. Figure 8A shows that initial stimulation of HBMEC with 1 nM thrombin effectively prevented [Ca2+]i changes in response to subsequent addition of 100 nM thrombin. However, pretreatment of the cells with thrombin did not prevent subsequent PAR1-AP-induced [Ca2+]i rise (Fig. 8, A and C). At the same time, pretreatment of cells with PAR1-AP partially desensitized subsequent Ca2+ responses to thrombin but it did not desensitize subsequent Ca2+ responses to PAR1-AP (Figs. 8B and 4B). These findings suggest that different types of desensitization of PAR1 occur in response to thrombin and PAR1-AP. Interestingly, we found that the trivalent cations La3+ (10–1,000 µM) and Gd3+ blocked thrombin-induced [Ca2+]i changes, whereas they did not significantly affect PAR1-AP-induced [Ca2+]i increase even at 5 mM La3+ (data not shown). Such a selective inhibition of thrombin-induced, but not PAR1-AP-induced, [Ca2+]i rise by La3+ suggests that La3+ may inhibit enzymatic activity of thrombin or interfere with a docking site of thrombin with the NH2 terminus of PAR1. Further studies using specific thrombin inhibitors such as D-phenylalanine-prolylarginine chloromethyl ketone (PPACK) or hirudin are needed to clarify this issue.



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Fig. 8. Pretreatment with low concentration of thrombin inactivated subsequent stimulation of HBMEC with higher concentrations of thrombin but not with PAR1-AP. A: HBMEC were sequentially challenged with 1 and 100 nM thrombin (Thr) and 100 µM PAR1-AP (P1). B and C: where indicated by horizontal lines, cells were stimulated with 100 µM PAR1-AP (P1) and 10 nM thrombin (Thr). Representative traces of single cells of 4 independent experiments are presented. Where indicated with arrow, 10 µM ATP was added.

 

Thrombin- and PAR1-AP-induced endothelial barrier dysfunction. One of the most prominent features of the brain endothelial cell barrier is a very low permeability to macromolecules and solutes compared with endothelial cells of nonbrain origin. Thrombin activation of endothelial cells of nonbrain origin is associated with a breakdown of endothelial barrier functions, gap formation, and increased permeability (8, 23). A decrease of transendothelial monolayer resistance is one of the signs of endothelial barrier dysfunction in response to thrombin stimulation (8, 23, 30, 39). We used a real-time TER measuring system to investigate the effects of thrombin and PAR1-AP on HBMEC. The addition of 10 nM thrombin caused a decrease in TER by 52% after 30 min, which was followed by a complete recovery after 90 min (Fig. 9). In contrast, stimulation of HBMEC with 100 µM PAR1-AP did not induce significant reduction of TER (Fig. 9). These results suggest that PAR stimulation by thrombin and PAR1-AP result in differential outcomes and also that PAR1-AP-induced Ca2+ influx is not sufficient to induce endothelial barrier dysfunction.



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Fig. 9. Effect of thrombin and PAR1-AP on transendothelial monolayer electrical resistance. Primary HBMEC were grown to confluence on gold microelectrode array. Cells were washed with HBSS, incubated with HBSS for 3 h, and challenged with 10 nM thrombin or 100 µM PAR1-AP where indicated with arrow. Representative traces are shown. Experiments were repeated 3 times.

 


    DISCUSSION
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In the present study, we report for the first time that primary HBMEC express all four known members of the PARs. Among four PARs, however, only PAR1 and PAR2 responded to corresponding PAR-AP by [Ca2+]i increase. These findings suggest that PAR3 and PAR4 may not be directly connected to calcium signaling in HBMEC. We showed in HBMEC that [Ca2+]i responses and activation of PAR1 differ between thrombin and PAR1-AP. For example, thrombin induced a dose-dependent rapid and transient [Ca2+]i increase, whereas the PAR1-AP SFFLRN triggered a dose-dependent rapid but prolonged rise in intracellular Ca2+. Thrombin- and PAR1-AP-induced [Ca2+]i increases were sensitive to thapsigargin and U-73122, supporting previous reports that PAR1 stimulation is associated with activation of PLC and Ca2+ discharge from endoplasmic reticulum (10). Subsequent studies revealed that PAR1 stimulation with PAR1-AP resulted in massive influx of extracellular Ca2+ through SOCCs, probably by a CCE mechanism (35, 47, 52). Removal of extracellular Ca2+ or preincubation with SKF-96365, a known blocker of SOCCs, eliminated the prolonged phase of PAR1-AP-induced [Ca2+]i rise (Fig. 7C). These findings suggest that activation of PAR1 with the untethered agonist peptide SFFLRN induced more severe depletion of intracellular Ca2+ stores and subsequent opening of SOCCs compared with thrombin-induced Ca2+ signaling by unmasked tethered agonist. Of interest, SFFLRN induced opening of SOCCs even in the absence of extracellular Ca2+. This was evident with readdition of extracellular Ca2+, which resulted in a strong increase of [Ca2+]i after SFFLRN stimulation, and this [Ca2+]i increase was blocked by SKF-95365, an inhibitor of SOCCs. In contrast, readdition of extracellular Ca2+ to the cells, which were previously stimulated by thrombin in the absence of extracellular Ca2+, did not produce any significant rise of [Ca2+]i.

Thrombin induces endothelial barrier dysfunction via two major pathways: one is calcium dependent and the other is RhoA dependent but calcium independent. The former pathway involves stimulation of myosin light chain (MLC) kinase through Ca2+/calmodulin, whereas RhoA stimulates Rho kinase (ROCK) and subsequent inhibition of MLC phosphatase (6, 7, 13, 15, 2931, 33, 39, 41, 49). Activation of both pathways results in formation of actin stress fibers and subsequent cell contraction. One of the indicators of endothelial barrier dysfunction is a drop in TER. By using a system of continuous monitoring of TER we found that thrombin decreased TER of HBMEC whereas PAR1-AP failed to induce a drop in this parameter. These findings suggest another potential difference in activation of PAR1 by tethered and untethered agonists in HBMEC. Furthermore, these observations suggest that PAR1-AP activation of the CCE mechanism is not necessary or sufficient to induce changes of cell shape/cell contraction and, hence, a drop in TER. These findings are in accordance with those of other studies that showed that stimulation of capacitative Ca2+ influx is not enough to induce cell contraction of cells (8, 53). In contrast, there are other reports in endothelial cells of nonbrain origin demonstrating that induction of CCE, for instance by thapsigargin-induced depletion of calcium stores, is associated with cell contraction and an increased permeability of the endothelial monolayer (40). These differences may be due to structural and functional differences between HBMEC and nonbrain endothelial cells. In addition, the heterogeneous responses to thrombin of various types of endothelial cells have been demonstrated, for example, for endothelial cells derived from lungs, coronary arteries, or umbilical cord veins (16, 19, 25). We showed in HBMEC that the PAR1-AP-induced Ca2+ influx involved SOCCs but did not cause significant drop in TER. It should also be mentioned that sensitivity of the endothelial cell barrier functions to store-operated calcium influx has been shown to be dependent on cell type (8, 25). For instance, receptor-independent activation of SOCCs, by preincubation of cells with thapsigargin, was sufficient to induce pulmonary macrovascular endothelial barrier dysfunction but not to affect pulmonary microvascular endothelial barrier function, suggesting that pulmonary micro- and macrovascular barrier functions are controlled by different mechanisms (8, 25). It is unclear whether a similar concept is relevant to thrombin action. Several groups have shown that thrombin-mediated permeability of pulmonary microvascular endothelial cells is calcium dependent (29, 31). However, in one study the role of CCE was not discriminated from the thrombin-induced general [Ca2+]i rise (31), and in another study only PAR1-AP, but not thrombin, induced store-operated calcium influx (29). Additionally, there are reports showing dramatic inhibition of thrombin-induced endothelial barrier dysfunction by Y-27632, an inhibitor of ROCK, which involves an essentially calcium-independent pathway (6, 7, 29, 31, 34). Whether particular mechanisms of thrombin-induced endothelial barrier dysfunction depend on the origin of the endothelial cells or whether both calcium-dependent and calcium-independent mechanisms are involved in thrombin-induced barrier dysfunction of HBMEC has yet to be clarified.

Thrombin can contribute to serious central nervous system disorders, such as hemorrhage and brain edema (17, 26, 51), if not tightly controlled. As a member of the seven-transmembrane spanning G protein-coupled receptor family, PAR1 can be activated indefinitely in the absence of a special mechanism of its desensitization. We speculate that interaction of thrombin with PAR1 in HBMEC is a highly regulated process, which may shut down immediately after initial interaction. For example, pretreatment of HBMEC with a low dose of thrombin completely prevented subsequent Ca2+ responses of the cells to much higher thrombin concentrations whereas PAR1-AP was still able to induce [Ca2+]i rise (Fig. 8). These data demonstrate the effectiveness of inactivation of thrombin-stimulated PAR1 in HBMEC. Such an ability of lower-dose thrombin to desensitize its receptor has also been shown for other types of endothelial cells (8, 23) and probably reflects a universal mechanism of desensitization of the activated receptor. Recent studies have provided some clues concerning how PAR1 is inactivated after thrombin stimulation (36, 46). It appears that G protein-coupled receptor kinases play a key role in inactivating PAR1 by the phosphorylation of agonist-occupied receptor (46). After phosphorylation the activated receptor apparently undergoes endocytosis and lysosomal degradation (36). However, our data as well as several other reports (36, 46) suggest that the desensitization of the activated PAR1 seems to depend on whether the receptor has been activated by thrombin and newly formed tethered ligand or by untethered activating peptide. Our demonstration of PAR1 desensitization to subsequent additions of thrombin but lack of desensitization to PAR1-AP after treatment of cells with lower-dose thrombin suggests that PAR1 is not immediately internalized and the extracellular binding sites remain available for PAR1-AP. Recent reports demonstrate that one of the key elements of thrombin desensitization of PAR1 is the binding of the special proteins {beta}-arrestins to the phosphorylated agonist-activated PAR1 (34). Such a binding of {beta}-arrestins effectively prevents subsequent stimulation with thrombin, however, without subsequent internalization of the activated receptor. These findings demonstrate that the fate of PAR1 differs from that of other G protein-coupled receptors, such as {beta}-adrenergic receptors (14, 37). We speculate that in HBMEC desensitization of PAR1 with thrombin, at least initially, involves {beta}-arrestins and that {beta}-arrestins do not affect sensitivity of PAR-1 to untethered PAR1-AP. Studies are in progress to examine such a possibility by using digital fluorescent image microscopy and immunofluorescent labeling of {beta}-arrestins and PAR1. Interestingly, extracellular mutations of PAR1 have been shown to exhibit profound differences in the abilities of thrombin and PAR1-AP to activate PAR1 receptor in Xenopus laevis oocytes transfected with PAR1 (5). For example, mutations strongly affected PAR1-AP-induced Ca2+ responses without significantly altering thrombin-induced Ca2+ changes or PAR1-AP binding to PAR1 receptor. These data suggest that conformational changes of PAR1 during agonist activation may also affect kinetics of PAR1-induced Ca2+ responses. It is possible that the observed differences of [Ca2+]i changes between thrombin and PAR1-AP stimulation of PAR1 in HBMEC may be related to different conformational changes induced by thrombin-unmasked tethered and PAR1-AP untethered peptides.

Another interesting finding in the present study is that La3+ inhibited thrombin-induced [Ca2+]i rise but failed to inhibit PAR1-AP-induced [Ca2+]i increase. The mechanism(s) of the inhibitory effect of La3+ on thrombin is not yet clear. Earlier reports showed that thrombin is a Na+-dependent serine protease (12, 50), and we can only speculate that La3+ probably competes with a Na+ binding site in thrombin and thus effectively inhibits its enzymatic activity. Another possibility is that La3+ may interfere with the docking site of thrombin with the NH2 terminus of PAR1.

In conclusion, we have shown for the first time that HBMEC express all four PAR receptors. Our studies of PAR1 signaling revealed that thrombin and PAR1-AP induce [Ca2+]i rises with different kinetics and mechanisms. The major difference was that, in contrast to thrombin, PAR1-AP stimulated opening of SOCCs, probably because of more severe depletion of the intracellular Ca2+ store. Compared with thrombin, the contrasting absence of desensitization of PAR1 responses to PAR1-AP after pretreatment of HBMEC with lower concentrations of thrombin suggest that PAR1 becomes quickly shut down to thrombin activation, however, without immediate internalization of the receptor. In addition, PAR1-AP failed to induce breakdown of the brain endothelial barrier as measured by TER whereas thrombin decreased TER in HBMEC. Further characterizations of the mechanisms involved in different responses to thrombin and PAR1-AP may help develop new strategies to control thrombin-mediated BBB dysfunctions.


    ACKNOWLEDGMENTS
 
GRANTS

This work was supported by National Institutes of Health Grants NS-26310, AI-47225, and HL-61651.


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. S. Kim, Division of Pediatric Infectious Diseases, Johns Hopkins Univ. School of Medicine, 600 N. Wolfe St., Park 256, Baltimore, MD 21287 (E-mail: kwangkim{at}jhmi.edu).

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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