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
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ABSTRACT |
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store-operated calcium influx; desensitization; transendothelial electrical resistance; digital imaging
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.
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EXPERIMENTAL PROCEDURES |
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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 300600 ·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 0.51% 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 -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, 2540 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
. After reaching maximal, steady-state readings of transendothelial resistance (
15,000
) 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 410 independent experiments. Probability values of P < 0.05 according to unpaired Student's t-test were considered significant.
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RESULTS |
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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, AC) 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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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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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 = 710; 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 = 4050 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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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+ (101,000 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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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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DISCUSSION |
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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 -arrestins to the phosphorylated agonist-activated PAR1 (34). Such a binding of
-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
-adrenergic receptors (14, 37). We speculate that in HBMEC desensitization of PAR1 with thrombin, at least initially, involves
-arrestins and that
-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
-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.
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ACKNOWLEDGMENTS |
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This work was supported by National Institutes of Health Grants NS-26310, AI-47225, and HL-61651.
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FOOTNOTES |
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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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