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Molecular and Cellular Biology, April 2006, p. 3231-3242, Vol. 26, No. 8
0270-7306/06/$08.00+0     doi:10.1128/MCB.26.8.3231-3242.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Clathrin Adaptor AP2 Regulates Thrombin Receptor Constitutive Internalization and Endothelial Cell Resensitization

May M. Paing,¹ Christopher A. Johnston,¹ David P. Siderovski,¹ and JoAnn Trejo^1,2*

Department of Pharmacology,¹ Department of Cell and Developmental Biology, School of Medicine, University of North Carolina, Chapel Hill, North Carolina 27599-7365²

Received 18 August 2005/ Returned for modification 30 September 2005/ Accepted 17 January 2006

Top Abstract Introduction Materials and Methods Results Discussion References

 
Protease-activated receptor 1 (PAR1), a G protein-coupled receptor for the coagulant protease thrombin, is irreversibly activated by proteolysis. Unactivated PAR1 cycles constitutively between the plasma membrane and intracellular stores, thereby providing a protected receptor pool that replenishes the cell surface after thrombin exposure and leads to rapid resensitization to thrombin signaling independent of de novo receptor synthesis. Here, we show that AP2, a clathrin adaptor, binds directly to a tyrosine-based motif in the cytoplasmic tail of PAR1 and is essential for constitutive receptor internalization and cellular recovery of thrombin signaling. Expression of a PAR1 tyrosine mutant or depletion of AP2 by RNA interference leads to significant inhibition of PAR1 constitutive internalization, loss of intracellular uncleaved PAR1, and failure of endothelial cells and other cell types to regain thrombin responsiveness. Our findings establish a novel role for AP2 in direct regulation of PAR1 trafficking, a process critically important to the temporal and spatial aspects of thrombin signaling.

Top Abstract Introduction Materials and Methods Results Discussion References

 
G protein-coupled receptors (GPCRs) comprise the largest family of receptors in the mammalian genome and elicit cellular responses to diverse extracellular stimuli (30). Intracellular trafficking of GPCRs controls temporal and spatial aspects of receptor signaling, including signal termination via removal of activated receptors from G-proteins and signaling effectors at the plasma membrane. Recent studies indicate that activated GPCRs can also signal internally at endocytic vesicles (1, 34). Once agonist dissociates from internalized receptor, GPCRs are then recycled back to the cell surface in a resensitized state competent to signal again. Trafficking of internalized GPCRs from endosomes to lysosomes with the consequent receptor degradation is also an important process that terminates receptor signaling (36, 37). The regulation of GPCR internalization, recycling, and lysosomal sorting involves specific interactions between receptor "sorting" motifs and endocytic adaptor proteins. However, the mechanisms that mediate trafficking of GPCRs through the endocytic pathway remain poorly defined.

Protease-activated receptor 1 (PAR1), prototype of a family of proteolytically activated GPCRs, is a receptor for the coagulant protease thrombin. PAR1 is the predominant mediator of thrombin signaling in human platelets, endothelial cells, fibroblasts, and smooth muscle cells and elicits a variety of cellular responses critical for normal vascular responses as well as cardiovascular disease processes (6, 21). PAR1 is activated by an unusual, irreversible proteolytic mechanism. Thrombin cleaves the extracellular amino terminus of the receptor, unmasking a new amino terminus that acts as a tethered ligand by binding intramolecularly to the receptor to trigger signaling (5, 38, 39). Synthetic peptides that mimic this newly formed amino-terminus can activate PAR1 independent of thrombin and receptor cleavage. The irreversible nature of proteolytic PAR1 activation, by generating a tethered ligand that cannot diffuse away, is distinct from the reversible activation of most GPCRs, raising the question, How do cells regulate thrombin signaling?

PAR1 trafficking is essential for the fidelity of thrombin signaling. In unstimulated fibroblasts and endothelial cells, PAR1 cycles constitutively between the cell surface and an intracellular compartment, forming a cytosolic receptor pool protected from thrombin cleavage and activation (13, 15, 18). Upon thrombin exposure, cell surface PAR1 is cleaved, activated and then internalized, sorted predominantly to lysosomes, and degraded (16, 36). Internalization and lysosomal sorting of irreversibly activated PAR1 are both critical for signal termination (36, 37). After thrombin is removed, uncleaved PAR1 moves from the intracellular protected pool to the cell surface. This replenishment of the cell surface with uncleaved PAR1 allows for rapid recovery of thrombin signaling independent of de novo receptor synthesis (13). However, the sorting motifs and endocytic adaptor proteins that specify the distinct trafficking behaviors of PAR1 are not known.

Arrestins are multifunctional adaptor proteins known to interact with the clathrin endocytic machinery to mediate GPCR internalization. We previously found that PAR1 internalization, although dependent on clathrin and dynamin, occurs independent of arrestins (4, 25, 35). Given this observation, and the presence of tyrosine-based motifs in the cytoplasmic tail of PAR1, we examined the function of the adaptor protein complex 2 (AP2). AP2 is a plasma membrane-localized clathrin adaptor composed of , ß2, µ2, and 2 adaptin subunits (3). The µ2 subunit binds directly to tyrosine-based sorting signals within the cytoplasmic regions of transmembrane proteins to facilitate internalization through clathrin-coated pits. Our studies here reveal that AP2 directly regulates PAR1 constitutive internalization and is essential for resensitization of endothelial cells and other cell types to thrombin signaling.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Reagents and antibodies. Human -thrombin was obtained from Enzyme Research Laboratories. The PAR1 agonist peptides SFLLRN and TFLLRNPNDK were synthesized as the carboxyl amide and purified by high-pressure liquid chromatography by the University of North Carolina Peptide Facility. N-terminal biotinylated peptides corresponding to the carboxy terminus of human PAR1 (amino acids 396 to 425) were synthesized and purified by high-pressure liquid chromatography by the Tufts University Core Facility (Boston, MA). Hirudin, cycloheximide, carbachol, isoproterenol, uridine triphosphate (UTP), and sucrose were purchased from Sigma. Calcium indicator dye Fura2-acetoxymethyl-ester (Fura2-AM), pluronic acid, 4-bromo A-23187 ionophore, transferrin-Alexa-488, and Alexa-488- and Alexa-594-conjugated goat anti-mouse antibodies were obtained from Molecular Probes. Monoclonal M1 anti-FLAG, ß-adaptin, ß-actin, and glutathione S-transferase (GST) antibodies were obtained from Sigma. Anti-AP50 (µ2), anti--adaptin, and anti-early endosome antigen (EEA1) monoclonal antibodies were from BD Biosciences. Polyclonal anti-His₆ antibody was obtained from Abcam Inc. Anti-ß-arrestin polyclonal antibody A1CT was generously provided by R. J. Lefkowitz (Duke University). A rabbit polyclonal anti-PAR1 antibody was generated against the amino-terminal peptide sequence YEPFWEDEEKNESGLTEYC, as previously described (17). Horseradish peroxidase-conjugated goat anti-mouse and anti-rabbit secondary antibodies were from Bio-Rad.

cDNAs and cell lines. A previously published cDNA encoding wild-type PAR1 containing an amino-terminal FLAG epitope was used to generate receptor mutants (18). Mutations were introduced using QuickChange site-directed mutagenesis (Stratagene) and confirmed by dideoxy sequencing. FLAG-tagged ß₂-adrenergic receptor cDNA was a gift from M. von Zastrow (University of California, San Francisco). A plasmid encoding green fluorescent protein (GFP)-tagged, dominant-negative (K44A) dynamin 2 was generously provided by M. McNiven (Mayo Clinic and Foundation). HeLa cells stably expressing FLAG-tagged PAR1 wild-type and mutants were generated and maintained as previously described (35). Human umbilical vein endothelial cells (HUVECs) were obtained from Clonetics and maintained according the manufacturer's instructions. HUVECs at early passages were used for all experiments.

siRNAs. HeLa cells were transiently transfected with 50 nM of nonspecific (NS) or µ2-specific small interfering RNAs (siRNAs) using Lipofectamine 2000 according to the manufacturer's instructions. Experiments were performed 60 h after transfections. HUVECs (1 x 10⁶) were electroporated with a 600 nM concentration of either NS or µ2 siRNA using technology developed by Amaxa, Inc., and experiments were carried out 48 h later. The µ2 siRNA targeting the mRNA sequence 5'-GTG GAT GCC TTT CGG GTC A-3' was previously described (20) and synthesized by Dharmacon, Inc. The NS siRNA 5'-CTA CGT CCA GGA GCG CAC C -3' was used as a negative control.

Internalization assays. Constitutive and agonist-induced PAR1 internalization were assessed using our previously published receptor-antibody uptake assay (26, 35).

Immunofluorescence confocal microscopy. HeLa cells or HUVECs were preincubated with rabbit polyclonal anti-PAR1 antibody for 1 h at 4°C to label the surface cohort, washed, and then left untreated or treated in the absence or presence of agonist for various times at 37°C. Cells were fixed and processed for confocal microscopy as we previously described (25, 40). Colocalization of ß2-adaptin was assessed by incubating permeabilized cells with anti-ß-adaptin antibody for 1 h at 25°C, followed by species-specific fluorophore-conjugated secondary antibodies, and then imaged by confocal microscopy. Colocalization of PAR1 with -adaptin was examined in fixed cells permeabilized with 0.2% Triton X-100 diluted in phosphate-buffered saline (PBS), after incubation with species-specific fluorophore-conjugated secondary antibodies using confocal microscopy. The extent of colocalization of PAR1 with either ß2-adaptin or -adaptin was quantitated by counting the number of PAR1-positive puncta that costained with ß2-adaptin or -adaptin, indicated by the yellow area in the merged image as previously described (40). The data are expressed as the percentage of PAR1-positive puncta that costained with either ß2-adaptin or -adaptin and represent the averages of many cell samples examined in independent experiments. To assess PAR1 colocalization with EEA1, permeabilized cells were incubated with anti-EEA1 antibody for 1 h at 25°C, washed, and then incubated with species-specific secondary fluorophore-conjugated antibodies and imaged by confocal microscopy. Transferrin uptake was assessed in serum-deprived HeLa cells by incubation with Alexa-488-conjugated transferrin for 1 h at 4°C; cells were washed and then warmed to 37°C for 10 min and internalized transferrin-Alexa-488 was imaged by confocal microscopy. Images were collected using a Fluoview 300 laser scanning confocal imaging system (Olympus) configured with an IX70 fluorescent microscope fitted with a PlanApo 60x oil objective. Fluorescent images, X-Y sections at 0.28 µm, were collected sequentially at 800 x 600 resolution with 2x optical zoom. The final composite image was created using Adobe Photoshop CS.

Immunoblotting. To detect µ2 expression, total cell lysates were resolved by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis, transferred to membranes, and immunoblotted with a monoclonal anti-AP50 (µ2) antibody. Blots were then stripped and reprobed with a monoclonal antiactin antibody. Immunoblots were developed with ECL-PLUS (Amersham) and imaged by autoradiography.

GST pull-down and surface plasmon resonance (SPR) binding assays. To generate GST fused to the PAR1 cytoplasmic (C)-tail domain, an EcoR I and BamH I insert containing a glycine/serine spacer (GSSG) and PAR1 C-tail residues 374 to 425 was ligated into compatible sites of pGEX2TK (Amersham). GST constructs were transformed into BL21(DE3) Escherichia coli, and fusion proteins were induced and purified using standard techniques previously described (41). A His₆-tagged µ2 expression plasmid (24), encompassing the carboxyl terminal amino acid residues 160 to 435 (provided by D. J. Owen, University of Cambridge, United Kingdom), was transformed into BL21(DE3)pLys-S E. coli. Six liters of culture was grown at 37°C to an optical density at 600 nm (OD₆₀₀) of 0.8 with progressive temperature reduction to 18°C prior to induction of His₆-µ2 expression with 50 µM IPTG (isopropyl-ß-D-thiogalactopyranoside) for 20 h at 18°C. The soluble cell lysate fraction was subjected to Ni²⁺-nitrilotriacetic acid (Ni-NTA) resin affinity chromatography, followed by size exclusion chromatography using established methods (41). Monodispersed fractions of His₆-µ2 protein were pooled and concentrated using a YM-10 centrifugal filter (Millipore) in a final buffer containing 20 mM HEPES, pH 7.5, 200 mM NaCl, 2 mM dithiothreitol, and 10% glycerol.

For GST pull-down binding assays, 20 µg of GST-PAR1 C-tail fusion protein or GST alone was immobilized on glutathione-Sepharose 4B beads and then incubated with 20 µg of His-tagged µ2 protein for 3 h at 4°C in binding buffer (10 mM Tris-HCl, pH 7.4, 5 mM EDTA, 0.2% Triton X-100). Binding reactions were then washed three times with binding buffer (without Triton X-100). Proteins that remained bound were eluted in 2x SDS gel loading buffer (100 mM Tris-HCl, pH 6.8, 10% SDS, 0.2% bromophenol blue, 1 mM dithiothreitol, and 20% glycerol), resolved by SDS-polyacrylamide gel electrophoresis, transferred to membranes, and immunoblotted with anti-His antibody or anti-GST antibody. Immunoblots were developed with ECL or ECL-PLUS (Amersham) and imaged by autoradiography.

SPR binding assays were performed at 25°C using a BIAcore 3000 in the University of North Carolina Pharmacology Protein Core. N-terminally biotinylated PAR1 C-tail peptides (diluted to 1 µg/ml in BIA running buffer [10 mM HEPES, pH 7.4, 150 mM NaCl, 3 mM EDTA, 0.005% NP-40]) were bound to separate flow cells of a streptavidin-coated biosensor chip (SA5; Biacore) to a surface density of 1,000 resonance units. Prior to injection, His₆-µ2 protein was diluted to desired concentrations in BIA running buffer. A total of 50 µl of His₆-µ2 protein was then simultaneously injected over flow cells at 10 µl/min, followed by a 200-s dissociation in BIA running buffer. All sensorgram curves were corrected for bulk buffer refractive index shifts and nonspecific binding by subtracting corresponding traces from a negative control blank surface. Following each injection, surfaces were regenerated with a 10-µl injection of 500 mM NaCl plus 20 mM NaOH at 20 µl/min. Binding curves and affinity calculations were conducted using BIAevaluation (version 3.0) and GraphPad Prism (version 4.0b).

Intracellular calcium measurements. Cells were grown on glass coverslips to a cell density of approximately 30 to 40% of confluence. Intracellular calcium was measured and quantified essentially as previously described (27). Cells loaded with 5 µM Fura2 were placed in a flowthrough chamber and superfused continuously with Hanks balanced salt solution in the presence or absence of agonists. Cells were exposed to alternating excitation wavelengths of 340 and 380 nm, and fluorescence emission at 510 nm was monitored using an integrating charge-coupled-device camera. The 340/380-nm fluorescence emission ratio was determined, and intracellular Ca²⁺ concentration was determined using the equation of Gyrnkiewicz et al. (12). The data were recorded and processed using InCyt IM2 digital imaging system (Intracellular Imaging Inc.).

Data analysis. Data were analyzed using Prism 4.0 software, and statistical significance was determined using InStat 3.0 (GraphPAD). Group comparisons were made using an unpaired Student's t test.

Top Abstract Introduction Materials and Methods Results Discussion References

 
PAR1 interacts directly with µ2 in vitro and colocalizes with endogenous AP2 in vivo. Using arrestin-deficient mouse embryonic fibroblasts, we previously found that PAR1 internalizes through a clathrin- and dynamin-dependent pathway independent of arrestin function (25). Arrestin-independent PAR1 internalization through clathrin-coated pits is also observed in HeLa cells and other cell types (4, 35) (see Fig. S1 in the supplemental material). To delineate the mechanism responsible for PAR1 internalization through clathrin-coated pits, we initially examined whether PAR1 could interact with the µ2 subunit of AP2. Purified, His₆-tagged µ2 protein was found to bind directly to a GST-PAR1 C-tail fusion protein absorbed to glutathione-Sepharose beads, but not to GST protein alone (Fig. 1A). We next determined whether PAR1 colocalizes with the clathrin adaptor AP2 in intact cells using confocal microscopy. HeLa cells stably expressing FLAG-tagged PAR1 were incubated with a polyclonal anti-PAR1 antibody at 4°C to ensure that only cell surface receptors would be labeled. Cells were then warmed to 37°C for 2.5 min in the absence of agonist; under these conditions unactivated PAR1 is recruited to clathrin-coated pits (35). Cells were then fixed and immunostained for PAR1 and the AP2 subunit ß2-adaptin. Approximately 30% of PAR1-positive puncta costained for ß2-adaptin (Fig. 1B), suggesting that the majority of constitutively internalized receptors recruited to clathrin-coated pits colocalize with AP2 in intact cells (Fig. 1B).

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FIG. 1. The µ2 subunit of AP2 interacts with the PAR1 C tail in vitro and PAR1 colocalizes with endogenous AP2 in intact cells. (A) GST-PAR1 C-tail fusion protein or GST protein alone absorbed to glutathione-Sepharose beads was incubated with purified His-tagged µ2. Bound proteins were eluted and immunoblotted with an anti-His antibody (upper panel). An aliquot of purified His-tagged µ2 representing 10% input is shown in the last lane. Membranes were stripped and reprobed with an anti-GST antibody to determine the total amount of GST-PAR1 C tail and GST protein loaded in various lanes (lower panel). (B) HeLa cells stably expressing FLAG-tagged PAR1 were preincubated with anti-PAR1 antibody for 1 h at 4°C such that only cell surface receptor-bound antibody. Cells were then incubated for 2.5 min at 37°C with no agonist, fixed, permeabilized, and immunostained for PAR1 (red) using a polyclonal anti-PAR1 antibody and for endogenous ß2-adaptin (green) using a monoclonal anti-ß2-adaptin antibody and imaged by confocal microscopy. Colocalization of PAR1 with AP2 is shown in yellow in the merged image. The image shown is representative of many cells examined in three independent experiments. Scale bar, 2 µm.

AP2 is essential for PAR1 constitutive internalization. To assess AP2 function in PAR1 trafficking, we used siRNA targeting the µ2 subunit to deplete HeLa cells of endogenous AP2 complex (20). Expression of µ2 was virtually abolished in cells transiently transfected with µ2 siRNA compared to cells transfected with a nonspecific control (Fig. 2A, inset). Cells transfected with µ2 siRNA also showed significant decreases in ß2-adaptin expression (Fig. 2C), consistent with a loss of endogenous AP2 complex (20). Inhibition of transferrin uptake in µ2 siRNA-transfected cells (see Fig. S2 in the supplemental material) further indicates that µ2 knockdown is sufficient to deplete HeLa cells of AP2 function, as the transferrin receptor is known to internalize constitutively and recycle through an AP2-, clathrin- and dynamin-dependent pathway (3).

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FIG. 2. PAR1 constitutive internalization, but not agonist-induced internalization, is inhibited in AP2-depleted HeLa cells. (A) HeLa cells stably expressing FLAG-tagged, wild-type PAR1 were transiently transfected with 50 nM siRNA targeted to either µ2 or nonspecific (ns) mRNA sequences. Surface FLAG-tagged PAR1 receptors were then labeled with the Ca2+-dependent M1 anti-FLAG antibody for 1 h at 4°C. Cells were then incubated (without agonist) for various times at 37°C to allow constitutive internalization of receptor-bound antibody. After incubations, cells were stripped of antibody remaining bound to the cell surface with PBS-0.04% EDTA and lysed, and internalized antibody was then quantified by enzyme-linked immunosorbent assay. Data (mean ± standard error of the mean; n = 3) are expressed as a percentage of initial cell surface receptor-bound antibody, defined as total antibody bound to cells at 0 min and not washed with PBS-EDTA. Similar results were obtained in at least three independent experiments. The immunoblot of equivalent amounts of cell lysates shown in the inset confirms the loss of µ2 protein in µ2 siRNA-transfected cells, whereas actin expression was unaffected. (B) PAR1-expressing HeLa cells transiently transfected with µ2 or ns siRNAs were labeled with antibody as described above and then incubated in the presence or absence of 50 µM SFLLRN agonist peptide for 10 min at 37°C. Internalized receptor-bound antibody was quantified as described above. The data (mean ± standard error of the mean; n = 3) shown are representative of at least three separate experiments. (C) PAR1-expressing HeLa cells transiently transfected with µ2 or ns siRNAs were fixed, permeabilized, and immunostained for ß2-adaptin expression and imaged by confocal microscopy. The differential interference contrast (DIC) image is the same field of µ2-transfected cells shown in the adjacent anti-ß2-adaptin fluorescence image. (D) PAR1-expressing HeLa cells transiently transfected with µ2 or ns siRNAs were either left untreated (0 min) or treated in the absence or presence of 50 µM SFLLRN for 10 min at 37°C. Cells were fixed, immunostained for PAR1, and imaged by confocal microscopy. The imaged cells are representative of many cells examined in three different experiments. Insets are magnifications of boxed areas. Scale bar, 10 µm.

To determine whether AP2 is necessary for PAR1 internalization, a stable HeLa cell line expressing FLAG-tagged PAR1 was labeled with anti-FLAG antibody and then incubated with or without agonist at 37°C for various times to allow internalization of receptor-bound antibody. In control siRNA-transfected cells not exposed to agonist, 20% of antibody initially bound to the cell surface was internalized at steady state (Fig. 2A and B), consistent with the extent of PAR1 constitutive internalization observed in other cell types (13, 32). In contrast, PAR1 constitutive internalization was inhibited significantly in µ2 siRNA-transfected cells (Fig. 2A and B), in spite of an increase in steady-state amounts of surface receptor in these cells (data not shown). These findings suggest that AP2 is necessary for PAR1 constitutive internalization. Exposure to the agonist peptide SFLLRN for 10 min caused substantial PAR1 internalization irrespective of siRNA treatment (Fig. 2B). Interestingly, in cells depleted of µ2, activated PAR1 internalization occurred through a clathrin- and dynamin-dependent pathway (see Fig. S3 in the supplemental material). These findings suggest that agonist-activated PAR1 internalization is independent of AP2 function.

Immunofluorescence studies of PAR1-expressing HeLa cells were consistent with an AP2-dependent regulation of receptor constitutive internalization. In control siRNA-transfected cells, a 10-min incubation at 37°C caused PAR1 to redistribute from the cell surface into endocytic vesicles (Fig. 2D), consistent with tonic cycling between the plasma membrane and an intracellular compartment. In contrast, cell surface PAR1 failed to internalize in unstimulated µ2 siRNA-transfected cells after 10 min at 37°C (Fig. 2D). Similar results were observed in cells transfected with a different µ2 siRNA targeting a distinct mRNA sequence (9) (data not shown). The addition of SFLLRN, however, induced significant accumulation of internalized PAR1 in both control- and µ2 siRNA-transfected cells (Fig. 2D), confirming that AP2 is not required for agonist-activated PAR1 internalization.

To determine whether AP2 regulates trafficking of endogenous receptor in native cells, we examined PAR1 internalization in HUVECs. HUVECs electroporated with µ2 siRNA showed significant loss of µ2 protein and ß2-adaptin expression compared to control siRNA-treated cells, suggesting that in these cells endogenous AP2 complex has been depleted (Fig. 3). A significant amount of internalized PAR1 was found in endocytic vesicles in control cells after 15 min at 37°C (Fig. 3B). By contrast, PAR1 failed to redistribute to endocytic vesicles in µ2 siRNA-transfected cells (Fig. 3B), suggesting an AP2-dependent regulation of PAR1 constitutive internalization in HUVECs. Incubation with the PAR1-specific agonist peptide TFLLRNPNDK caused a marked increase in receptor-containing endocytic vesicles in both control- and µ2 siRNA-transfected cells (Fig. 3C), indicating that activated PAR1 internalizes independent of AP2. Together these findings reveal a novel function for AP2 in regulating PAR1 constitutive internalization.

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FIG. 3. AP2 is essential for endogenous PAR1 constitutive internalization, but not agonist-induced internalization, in HUVECs. (A) HUVECs were electroporated with µ2 or nonspecific (ns) siRNAs. Cell lysates were prepared and immunoblotted (IB) for µ2 expression or actin expression. (B and C) HUVECs electroporated with µ2 or ns siRNAs were incubated with anti-PAR1 antibody for 1 h at 4°C; under these conditions only receptors on the cell surface bound antibody. Cells were washed and then incubated in the absence or presence of 100 µM TFLLRNPNDK agonist peptide for 15 min at 37°C. Cells were fixed, immunostained for PAR1 and ß2-adaptin expression, and imaged by confocal microscopy. These images are representative of many cells examined in at least three independent experiments. The insets are magnifications of boxed areas. Scale bar, 10 µm.

A distal tyrosine-based motif within PAR1 is required for µ2 binding and constitutive internalization. The µ2 subunit of AP2 binds directly to tyrosine-based sequences in cytoplasmic domains of transmembrane proteins and thereby facilitates recruitment to clathrin-coated pits at the cell surface (3). A search of the PAR1 sequence for such sorting signals revealed two highly conserved sequences in the C-tail region conforming to the YXXØ motif (where Y denotes tyrosine, X is any amino acid, and Ø is a bulky hydrophobic amino acid) (Fig. 4A). The interaction of µ2 with tyrosine-based sequences is strictly dependent on the tyrosine and bulky hydrophobic residue positions; however, residues flanking the motif also contribute to µ2 binding (3). To test the role of each tyrosine-based motif in PAR1 trafficking, we generated receptor mutants in which critical positions (Y³⁸³ and L³⁸⁶ or Y⁴²⁰ and L⁴²³) were converted to alanines (Fig. 4A). The L⁴²⁴ residue flanking the distal tyrosine-based motif was also mutated to alanine to avoid the possibility of compensatory effects on µ2 binding. In wild-type PAR1-expressing HeLa cells, 10 to 20% of antibody-labeled cell surface receptor was constitutively internalized at steady state (Fig. 4B and C). The PAR1 A³⁸³SIA³⁸⁶ mutant exhibited levels of constitutive and agonist-induced internalization similar to wild-type (Fig. 4B and C), indicating that the proximal tyrosine-based motif is not essential for either process. In striking contrast, however, PAR1 A⁴²⁰KKA⁴²³A⁴²⁴ mutants failed to constitutively internalize, suggesting that the distal tyrosine-based motif functions in PAR1 tonic cycling (Fig. 4B and C). A similar defect in constitutive internalization was observed upon mutation of only the distal tyrosine (Y⁴²⁰) to alanine, whereas mutation of the L⁴²³L⁴²⁴ residues had no significant effect (Fig. 4D), indicating that the critical Y⁴²⁰ is essential for function. However, when activated, the PAR1 A⁴²⁰KKA⁴²³A⁴²⁴ mutant internalized similar to wild-type receptor (Fig. 4C), indicating that the distal tyrosine-based motif is not essential for agonist-activated receptor internalization.

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FIG. 4. A PAR1 cytoplasmic tail distal tyrosine-based motif regulates constitutive but not agonist-induced receptor internalization. (A) PAR1 cytoplasmic tail amino acid sequence is shown, and the positions of the tyrosine-based motifs are indicated. Superscripted numbers indicate the critical tyrosines and leucines that were mutated to alanines. The asterisk indicates the end of the protein sequence. (B) HeLa cells stably expressing similar amounts of FLAG-tagged, wild-type PAR1 or receptor mutants were labeled with antibody and incubated (without agonist) for various times at 37°C, and internalized receptor was quantified as described above. The data (mean ± standard error of the mean; n = 3) shown are expressed as a percentage of total cell surface receptor-bound antibody at 0 min and are representative of at least three separate experiments. (C) HeLa cells stably expressing wild-type PAR1 or receptor mutants were labeled with antibody and incubated in the presence or absence of 50 µM SFLLRN agonist peptide for 10 min at 37°C, and the amount of receptor internalization was quantitated as described above. Data are representative of at least three independent experiments. (D) HeLa cells expressing comparable amounts of FLAG-tagged PAR1 wild-type, LL423/424AA, or Y420A mutants were labeled with antibody, washed, incubated in the presence or absence of 50 µM SFLLRN agonist peptide for 30 min at 37°C, and then quantitated. The data (mean ± standard error of the mean; n = 3) shown are representative of at least three independent experiments. A significant difference in constitutive endocytosis of the Y420A mutant compared to wild-type receptor was detected (*, P < 0.05), whereas no significant difference in constitutive endocytosis of the LL423/424AA mutant compared to wild-type receptor was detected. Statistical analysis was determined using an unpaired Student's t test. WT, wild-type; ASIA, A383SIA386; AKKAA, A420KKA423A424.

Immunofluorescence confocal microscopy was performed to confirm the function of tyrosine-based motifs in receptor trafficking. In untreated control cells, wild-type PAR1 and YXX Ø mutants were each found predominantly at the cell surface and did not colocalize with EEA1, a specific marker of early endosomes (Fig. 5a to d). After 10 min at 37°C, wild-type PAR1 and A³⁸³SIA³⁸⁶ mutant moved from the cell surface to endocytic vesicles and markedly colocalized with EEA1, suggesting that constitutively internalized receptor transits through an early endosomal compartment (Fig. 5a' and b'). Addition of SFLLRN caused an even greater increase in wild-type PAR1 and A³⁸³SIA³⁸⁶ mutant internalization (Fig. 5a" and b"). In contrast, A⁴²⁰KKA⁴²³A⁴²⁴ mutant PAR1 failed to redistribute to EEA1-positive endosomes after 10 min at 37°C (Fig. 5c' and d'), consistent with a lack of receptor constitutive internalization. However, agonist treatment induced a significant amount of A⁴²⁰KKA⁴²³A⁴²⁴ mutant internalization comparable to that of wild-type receptor (Fig. 5c" and d").

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FIG. 5. Immunofluorescence reveals a critical role for the PAR1 cytoplasmic tail distal tyrosine-based motif in constitutive but not agonist-induced internalization. HeLa cells expressing comparable amounts of wild-type (WT) PAR1 or receptor mutants were either left untreated (a to d; 0 min) or incubated in the absence (a' to d'; 10 min) or presence (a" to d"; +SFLLRN 10 min) of 50 µM SFLLRN agonist peptide for 10 min at 37°C. Cells were fixed and coimmunostained for PAR1 (green), using a polyclonal anti-PAR1 antibody, and for the early endosome marker EEA1 (red), using a monoclonal anti-EEA1 antibody, and imaged by confocal microscopy. The insets are magnifications of boxed areas. The imaged cells are representative of many cells examined in at least three independent experiments. Scale bar, 10 µm. ASIA, A383SIA386; AKKAA, A420KKA423A424.

We next determined whether the distal tyrosine-based motif (Y⁴²⁰KKL⁴²³) was important for PAR1 colocalization with endogenous AP2 in cells. HeLa cells expressing wild-type PAR1 or A⁴²⁰KKA⁴²³A⁴²⁴ mutant were labeled with a polyclonal anti-PAR1 antibody at 4°C so that only the surface receptor cohort binds antibody. PAR1 recruitment to clathrin-coated pits was then initiated by warming cells to 37°C for 2.5 min (35). Cells were fixed and immunostained for PAR1 and -adaptin, and colocalization was assessed by confocal microscopy. Approximately 29% of wild-type PAR1 localized to distinct puncta that costained for the AP2 subunit -adaptin (Fig. 6a). In contrast, A⁴²⁰KKA⁴²³A⁴²⁴ mutant remained predominantly in puncta that showed minimal colocalization with -adaptin (Fig. 6b); only 8% of A⁴²⁰KKA⁴²³A⁴²⁴ mutant-positive puncta costained for the -adaptin subunit. Thus, mutation of the distal tyrosine-based motif disrupts PAR1 colocalization with endogenous AP2 in intact cells, consistent with the failure of mutant receptor to undergo constitutive internalization.

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FIG. 6. The distal tyrosine motif is important for PAR1 colocalization with AP2 in intact cells. HeLa cells expressing PAR1 wild-type (WT) or the distal tyrosine motif mutant A420KKA423A424 (AKKAA) were prelabeled with anti-PAR1 antibody, washed, and then warmed to 37°C for 2.5 min to initiate PAR1 recruitment to clathrin-coated pits. Cells were fixed and coimmunostained for PAR1, (green) using a polyclonal anti-PAR1 antibody, or for the AP2 subunit -adaptin (red), using a monoclonal anti--adaptin antibody, and imaged by confocal microscopy. Colocalization of PAR1 with AP2 is revealed by the yellow areas in the merged image. The insets are magnifications of boxed areas. The image shown is representative of many cells examined in three different experiments. Scale bar, 10 µm.

These findings from cellular expression of PAR1 YXXØ mutants suggest that the distal tyrosine-based motif Y⁴²⁰KKL⁴²³ is critical to PAR1 constitutive internalization and colocalization with AP2 in intact cells. We therefore tested whether this distal motif was necessary for the observed direct binding of µ2 protein to the PAR1 C-tail (Fig. 1A). Recombinant His₆-tagged µ2 protein, purified from E. coli expression by nickel-NTA and size exclusion chromatography (Fig. 7A), was injected over streptavidin-coated SPR biosensors containing immobilized biotinylated PAR1 C-tail peptides (Fig. 7B and C). The dissociation constant (K_D) for the binding of µ2 to the wild-type PAR1 C-tail was 7.5 ± 1.3 µM. In contrast, no detectable binding was observed to the A⁴²⁰KKA⁴²³A⁴²⁴ mutant PAR1 C-tail peptide at any concentration of µ2 protein tested (Fig. 7C), indicating that the distal YXXØ motif is indeed required for direct µ2 association.

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FIG. 7. The distal tyrosine-based motif is necessary for direct association between the PAR1 cytoplasmic tail and purified recombinant µ2 protein. (A) His6-tagged µ2 protein was expressed in E. coli by IPTG induction, and soluble, monodispersed protein was purified by sequential application of Ni-NTA resin and size exclusion chromatography. (B) Representative binding curves were generated by injecting 15 µM purified His6-tagged µ2 protein over immobilized PAR1 C-tail peptide surfaces, followed by subtracting corresponding traces from a blank negative control surface. (C) Increasing concentrations of His6-tagged µ2 protein (0.01 to 50 µM) were injected separately over peptide biosensor surfaces, and binding curves were generated as described in panel B. The resulting maximal resonance units obtained during the analyte association phase were then plotted versus analyte concentration to generate a one-site binding curve, indicating a dissociation constant (KD) of 7.5 ± 1.3 µM for wild-type PAR1 C tail. No detectable binding was observed for the PAR1 A420KKA423A424 (AKKAA) mutant peptide surface, even at the saturating concentration of 50 µM.

Recovery of cellular thrombin signaling is dependent on AP2-regulated PAR1 constitutive internalization. Constitutive cycling of PAR1 between the plasma membrane and an intracellular compartment results in a protected pool of uncleaved receptors that repopulate the cell surface following thrombin exposure without de novo receptor synthesis (13). This process is critical for rapid cellular resensitization to thrombin signaling. We therefore assessed the role of AP2 and tyrosine-based motifs in the recovery of cellular thrombin responsiveness by measuring thrombin-stimulated increases in intracellular Ca²⁺ after an initial thrombin exposure in the presence of cycloheximide, a protein synthesis inhibitor. Adding a saturating concentration of thrombin resulted in cleavage and activation of all surface-bound wild-type or mutant PAR1 expressed in HeLa cells (data not shown). In wild-type PAR1-expressing cells, an initial exposure to thrombin induced a transient increase in cytosolic Ca²⁺ (Fig. 8A), comparable to that observed in other cell types (13). Cells were immediately refractory to thrombin. After thrombin removal, cells were allowed to recover for 20 min in the presence of hirudin (a thrombin inhibitor) and cycloheximide and then exposed to thrombin again, which elicited a second rise in intracellular Ca²⁺ (Fig. 8A). This time course of recovery is similar to that observed with PAR1-expressing fibroblasts (13). Cells expressing the PAR1 A³⁸³SIA³⁸⁶ mutant also exhibited recovery of thrombin signaling after an initial thrombin exposure (Fig. 8B). These findings are consistent with the idea that wild-type PAR1 and A³⁸³SIA³⁸⁶ mutant undergo constitutive internalization, generating an internal pool of protected receptors that recycles back to the cell surface (13, 26). Cells expressing the PAR1 A⁴²⁰KKA⁴²³A⁴²⁴ mutant displayed an initial thrombin-stimulated increase in cytosolic Ca²⁺ comparable to wild-type receptor; however, these cells failed to elicit a second response to thrombin (Fig. 8C and D). The ability of PAR1 A⁴²⁰KKA⁴²³A⁴²⁴ mutant-expressing cells to respond to carbachol, an agonist for endogenous muscarinic acetylcholine receptors, indicates that the cells were not globally defective in GPCR-stimulated Ca²⁺ mobilization after initial thrombin exposure (Fig. 8E and F). Thus, the failure of PAR1 A⁴²⁰KKA⁴²³A⁴²⁴ mutant-expressing cells to regain thrombin signaling appears to be linked to the lack of an internal pool of uncleaved receptors that replenish the cell surface and permit recovery of thrombin responsiveness.

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FIG. 8. PAR1 mutants defective in constitutive internalization fail to regain thrombin responsiveness. HeLa cells stably expressing similar amounts of wild-type (WT) PAR1 or receptor mutants grown on coverslips were pretreated with 10 µM cycloheximide for 90 min to block de novo protein synthesis, loaded with Fura2-AM, and then stimulated with thrombin (-Th) as indicated. The Ca2+ responses (i.e., [Ca2+]i, intracellular Ca2+ concentration) were quantified as described in Materials and Methods. Cells were first exposed to 30 nM thrombin for 10 min at 37°C, and the traces shown represent Ca2+ responses of at least 10 cells from a single field, all of which responded to thrombin. The presence of agonists and time of exposure are indicated by the bars below and above the traces. Between stimulations, cells were washed with medium containing the thrombin inhibitor hirudin (0.5 U/ml) and allowed to recover for 20 min at 37°C. (A and B) Wild-type (WT) PAR1 and PAR1 A383SIA386 mutant-expressing cells responded to a second 10 nM thrombin stimulus as well as to 2 µM A23187, a calcium ionophore. Similar results were observed in three independent experiments. (C to F) HeLa cells expressing the PAR1 A420KKA423A424 mutant failed to elicit a second response to 10 nM thrombin but retained the capacity to respond to 500 µM carbachol (CARB), an agonist for endogenous muscarinic acetylcholine GPCRs, or to the calcium ionophore A23187 (2 µM). Similar findings were observed in three separate experiments. ASIA, A383SIA386; AKKAA, A420KKA423A424.

We next examined whether AP2 regulation of PAR1 constitutive internalization affected cellular resensitization to thrombin signaling. In PAR1-expressing HeLa cells transfected with control siRNA, thrombin elicited an initial increase in cytosolic Ca²⁺, and after recovery virtually all cells regained thrombin responsiveness (Fig. 9A and C). An initial Ca²⁺ response to thrombin was also observed in 90% of µ2 siRNA-transfected cells; however, after recovery only 20% of cells were capable of eliciting a second response to thrombin (Fig. 9B and C). In contrast, carbachol-induced calcium mobilization was maintained in the majority of µ2 siRNA-transfected cells following thrombin exposure (Fig. 9D), indicating that cells were capable of responding to other GPCR agonists.

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FIG. 9. AP2 is essential for restoring thrombin signaling in HUVECs and other cell types. PAR1-expressing HeLa cells were transiently transfected with 50 nM µ2 or NS siRNA, and HUVECs were electroporated with 600 nM µ2 or nonspecific (ns) siRNAs as described in Materials and Methods. Cells were grown on coverslips, pretreated with 10 µM cycloheximide, loaded with Fura2-AM, and then exposed to thrombin (-Th) as indicated. The intracellular Ca2+ response traces from individual cells are shown in various panels, whereas the responses from an average of 20 to 30 cells imaged from three separate experiments are shown as bar graphs in panels C, D, G, and H. Cells were first exposed to 30 nM thrombin for 10 min at 37°C, and 80% of these cells responded to thrombin. Cells were washed with medium containing hirudin (0.5 U/ml) and then exposed to 10 nM thrombin again. In control ns siRNA-treated cells, thrombin elicited a second Ca2+ response in virtually the same percentage of cells that were initially responsive to thrombin. In µ2 siRNA-treated cells, only 1 to 2 cells out of 20 cells were responsive to thrombin after an initial thrombin exposure (panels C and G). The difference between the number of cells eliciting -thrombin-induced calcium response in µ2 siRNA-treated cells compared to ns siRNA control-treated cells was significant (**, P < 0.01; *, P < 0.05) as determined using an unpaired Student's t test. HeLa cells and HUVECs retained the capacity to respond to 500 µM carbachol (CARB) or 100 µM UTP, agonists for endogenously expressed muscarinic and purinergic GPCRs, respectively (panels D and H), and to the calcium ionophore A23187 (2 µM). Similar results were observed in at least three independent experiments.

Resensitization of HUVECs to thrombin signaling was also assessed. Nearly 80% of HUVECs electroporated with control siRNA were initially responsive to thrombin and immediately refractory to a second thrombin exposure, but after a 20-min recovery period, almost all cells (75%) again elicited a thrombin response (Fig. 9E and G). These findings are consistent with the recovery of thrombin responsiveness previously observed in these cells (13). In contrast, however, only 10% of HUVECs with siRNA-mediated µ2 knockdown regained thrombin responsiveness following initial thrombin exposure and a recovery period (Fig. 9F and G). Signaling by UTP, an agonist for endogenous P2Y₂ and/or P2Y₄ GPCRs, was unperturbed, indicating that µ2 siRNA-electroporated cells are responsive in general to GPCR activation after an initial thrombin exposure (Fig. 9H). Taken together, these findings strongly suggest that AP2-dependent regulation of PAR1 constitutive internalization is essential for recovery of cellular responses to thrombin.

Top Abstract Introduction Materials and Methods Results Discussion References

 
In the present study, we have defined a hitherto unknown role for the clathrin adaptor AP2 in regulating PAR1 constitutive internalization and restoring thrombin responsiveness to endothelial cells and other cell types. The µ2 subunit of AP2 directly interacts with a tyrosine-based motif present at the extreme carboxy terminus of PAR1. This tyrosine-based motif and AP2 function are each critical for maintaining an intracellular store of protected receptors that replenish the cell surface with uncleaved PAR1 after thrombin exposure, leading to recovery of thrombin signaling independent of de novo receptor synthesis. Interestingly, internalization of agonist-activated PAR1 through clathrin-coated pits is independent of AP2, suggesting that constitutive and agonist-induced receptor internalization are specified by distinct molecular machinery. These findings demonstrate a direct role for AP2 in regulation of PAR1 signaling and trafficking.

Most GPCRs, including the well-characterized archetype of the ß₂-adrenergic receptor, are rapidly phosphorylated following activation, bind arrestins, and are then recruited to a clathrin- and dynamin-dependent pathway for internalization from the plasma membrane (33). Arrestins facilitate GPCR internalization by binding directly to the clathrin heavy chain and the ß2-adaptin subunit of AP2 (8, 10). We have shown in multiple cell types that arrestins are not essential for PAR1 constitutive or agonist-induced internalization (4, 25). In contrast, we report here that the µ2 subunit of AP2 directly binds to a tyrosine-based motif in the cytoplasmic tail of PAR1 to mediate constitutive internalization. Mutation of PAR1 distal tyrosine-based motif or depleting cells of AP2 by RNA interference resulted in significant inhibition of receptor constitutive internalization. Moreover, the PAR1 distal tyrosine-based motif was also important for receptor colocalization with endogenous AP2 in intact cells and critical for direct µ2 binding in vitro. The low-affinity binding of PAR1 with the µ2 subunit of AP2 as observed in our studies (K_D, 7.5 µM) is typical of the moderate to weak interaction of µ2 with proteins bearing tyrosine-based motifs previously reported (22, 31) and may have precluded our ability to detect PAR1 interaction with AP2 using cell lysate-based approaches of coimmunoprecipitation or pull-downs. Such low-affinity protein-protein interactions are biologically relevant and, in fact, are required to allow for the dynamic assembly and disassembly of the protein network that drives clathrin-coated vesicle budding at the plasma membrane (23). A high-affinity interaction between PAR1 and µ2 would be detrimental, resulting in significant internalization of the receptor at steady state and rendering it unavailable to its extracellular stimuli.

In addition to PAR1, we found that several other human GPCRs contain sequences conforming to the canonical YXXØ motif within their cytoplasmic tails (see Fig. S4 in the supplemental material), suggesting that AP2 might function in trafficking of other GPCRs. Indeed, the µ2 subunit of AP2 was previously shown to directly interact with the C tail of the _1b-adrenergic receptor (7). However, in this case, µ2 binds to an unusual stretch of eight arginine residues (rather than a canonical YXXØ motif) within the C-tail of the _1b-adrenergic receptor. This study also reported that an _lb-adrenergic receptor mutant, in which the arginine stretch was deleted, failed to bind µ2 in vitro, and displayed defects in agonist-induced endocytosis, suggesting that this region (and perhaps AP2 binding) is important for activated _lb-adrenergic receptor internalization. The constitutively active human viral chemokine US28 GPCR has also been reported to internalize through a clathrin-dependent pathway involving AP2 and not arrestins (9). However, whether AP2 directly binds to a YXXØ motif present in the C-tail region of US28 receptor remains to be determined. Together, these findings suggest that the clathrin adaptor AP2 functions in distinct modes of GPCR trafficking, including constitutive internalization of unactivated PAR1, activated _1b-adrenergic receptor internalization, and uptake of the constitutively active viral chemokine US28 receptor.

Our study also indicates that the trafficking behaviors of unactivated versus activated PAR1 are regulated by distinct endocytic machinery. PAR1 constitutive internalization was virtually abolished in cells depleted of AP2, whereas agonist-activated receptor internalization remained intact (Fig. 2 and 3). Activated PAR1 internalized via a clathrin- and dynamin-dependent pathway in AP2 depleted cells, excluding the possibility that the receptor uses an alternate non-clathrin endocytic route (see Fig. S3 in the supplemental material). Our previous studies suggest that internalization of agonist-activated PAR1 is critically dependent on phosphorylation of the C-tail region (25, 32). Activated PAR1 phosphorylation might induce a conformational change within the C-tail region to expose an endocytic-sorting motif and/or to promote binding of clathrin adaptor proteins. We previously showed that mutation of the proximal tyrosine-based motif partially inhibits the endocytosis of activated PAR1 that is only evident after prolonged agonist exposure (26). However, AP2 does not appear to be involved in this process since, in the current studies, we find that AP2 is not essential for activated PAR1 internalization (Fig. 2 and 3). Thus, whether other clathrin adaptor proteins such as epsin or CALM bind directly to this motif or elsewhere to PAR1 intracytosolic domains to regulate AP2-independent internalization of activated PAR1 through clathrin-coated pits is not known. Internalization of activated PAR1 may be additionally dependent on ubiquitination or deubiquitination processes. The transient modification of GPCRs and/or associated proteins by ubiquitination is known to facilitate trafficking through the endocytic pathway (14).

The proteolytic activation of PAR1 is unique among GPCRs, and distinct intracellular pathways have evolved to dispose of irreversibly activated receptors and to replenish the cell surface with uncleaved receptors after thrombin exposure. Our findings strongly suggest that AP2-dependent constitutive internalization of PAR1 is critical for resensitizing cells to thrombin signaling. In human platelets, which presumably respond to thrombin only once, the majority of PAR1 is retained on the cell surface and fails to internalize (19). Thus, platelets lack an internal pool of protected receptors as recovery of thrombin responsiveness is not a physiological requirement for this cell type. In contrast, endothelial cells and fibroblasts are exposed to thrombin repeatedly and need to recover thrombin signaling in a timely manner. This appears to be accomplished by the movement of uncleaved PAR1 from an intracellular compartment to the cell surface to permit rapid recovery of thrombin signaling independent of de novo receptor synthesis (13). Evidence presented here supports this model. In cells expressing PAR1 tyrosine mutants or in cells depleted of AP2 by RNA interference, uncleaved receptor fails to constitutively internalize and accumulate in an intracellular store. Consequently, PAR1 is not protected from thrombin cleavage and cells are unable to regain thrombin responsiveness. However, cells retained the capacity to elicit calcium responses to other endogenous GPCR agonists, indicating that receptor activation is not globally disrupted. Several other GPCRs including PAR2 and the thromboxane A2 receptor ß isoform have also been reported to constitutively internalize, resulting in an intracellular store of unactivated receptors (2, 28). A noncanonical YX₃Ø motif is present in the cytoplasmic tail of thromboxane A2 receptor (29), whereas PAR2 lacks such tyrosine-based sequences. The mechanism mediating constitutive internalization of these receptors as well as its physiological relevance remains poorly defined.

Our studies provide the first insight into the molecular mechanisms responsible for PAR1 constitutive internalization and cellular resensitization to thrombin signaling. The proteolytic nature of PAR1 activation is distinct from most reversibly activated GPCRs. Consequently, endothelial cells and other cell types, which need to respond to thrombin repeatedly over time, have evolved specialized mechanisms to permit rapid recovery or maintenance of thrombin responsiveness. Thrombin and PAR1 are important mediators of endothelial cell activation and migration during vascular development and in response to vascular injury (11, 42). Cell migration requires repeated sampling of the local environment in order to detect changes in agonist concentrations and gradients. It will be important to determine whether disruption of PAR1 tonic cycling and failure of endothelial cells to rapidly recover thrombin signaling are critical for endothelial cell activation and/or migration in vivo.

 
We thank T. K. Harden and R. A. Nicholas for comments and helpful discussions and David J. Owen and Robert J. Lefkowitz for generously providing reagents.

This work was supported by National Institutes of Health grants HL067697 and HL073328 (to J.T.) and GM065533 (to D.P.S.). M.M.P. and C.A.J. were each supported by American Heart Association Predoctoral Fellowships.

 
* Corresponding author. Mailing address: Department of Pharmacology, University of North Carolina at Chapel Hill, 1106 Mary Ellen Jones Bldg., Chapel Hill, NC 27599-7365. Phone: (919) 843-7691. Fax: (919) 966-5640. E-mail: joann_trejo{at}med.unc.edu.

Supplemental material for this article may be found at http://mcb.asm.org/.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Molecular and Cellular Biology, April 2006, p. 3231-3242, Vol. 26, No. 8
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