Receptor/ß-Arrestin Complex Formation and the Differential Trafficking and Resensitization of ß2-Adrenergic and Angiotensin II Type 1A Receptors

Pieter H. Anborgh, Jennifer L. Seachrist, Lianne B. Dale and Stephen S. G. Ferguson

The John P. Robarts Research Institute and Departments of Physiology (J.L.S., S.S.G.F.) and Pharmacology and Toxicology (L.B.D., S.S.G.F.) University of Western Ontario London, Ontario, Canada N6A 5K8


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
ß-Arrestins target G protein-coupled receptors (GPCRs) for endocytosis via clathrin-coated vesicles. ß-Arrestins also become detectable on endocytic vesicles in response to angiotensin II type 1A receptor (AT1AR), but not ß2-adrenergic receptor (ß2AR), activation. The carboxyl-terminal tails of these receptors contribute directly to this phenotype, since a ß2AR bearing the AT1AR tail acquired the capacity to stimulate ß-arrestin redistribution to endosomes, whereas this property was lost for an AT1AR bearing the ß2AR tail. Using ß2AR/AT1AR chimeras, we tested whether the ß2AR and AT1AR carboxyl-terminal tails, in part via their association with ß-arrestins, might regulate differences in the intracellular trafficking and resensitization patterns of these receptors. In the present study, we find that ß-arrestin formed a stable complex with the AT1AR tail in endocytic vesicles and that the internalization of this complex was dynamin dependent. Internalization of the ß2AR chimera bearing the AT1AR tail was observed in the absence of agonist and was inhibited by a dominant-negative ß-arrestin1 mutant. Agonist-independent AT1AR internalization was also observed after ß-arrestin2 overexpression. After internalization, the ß2AR, but not the AT1AR, was dephosphorylated and recycled back to the cell surface. However, the AT1AR tail prevented ß2AR dephosphorylation and recycling. In contrast, although the ß2AR-tail promoted AT1AR recycling, the chimeric receptor remained both phosphorylated and desensitized, suggesting that receptor dephosphorylation is not a property common to all receptors. In summary, we show that the carboxyl-terminal tails of GPCRs not only contribute to regulating the patterns of receptor desensitization, but also modulate receptor intracellular trafficking and resensitization patterns.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
G protein-coupled receptor (GPCR) responsiveness to agonist wanes rapidly with time, a process termed desensitization. GPCR desensitization occurs as the consequence of rapid receptor phosphorylation by both second messenger-dependent protein kinases and G protein-coupled receptor kinases (GRKs) (reviewed in Refs. 1, 2, 3). GRK-mediated phosphorylation promotes the binding of arrestin proteins that sterically uncouple receptors from their cognate heterotrimeric G protein (4, 5). It has been known for some time that desensitized GPCRs also internalize in clathrin-coated vesicles (6, 7). More recently, both ß-arrestin1 and ß-arrestin2 were shown to mediate the internalization of receptors via clathrin-coated pits (8, 9, 10). ß-Arrestin-dependent receptor internalization via clathrin-coated vesicles is the consequence of ß-arrestin protein interactions with both the ß-adaptin subunit of the AP2 adaptor complex and clathrin (11, 12). In the case of the ß2-adrenergic (ß2AR)1 and neurokinin 1 receptors, internalization to the endosomal compartment results in ligand dissociation followed by receptor dephosphorylation (13, 14, 15, 16, 17). Subsequently, these receptors are recycled back to the plasma membrane surface as fully functional receptors (13, 14, 15). This has led to the suggestion that GPCR internalization represents the predominant mechanism underlying receptor resensitization. However, this pattern is not universal, since not all GPCRs recycle back to the cell surface after their internalization. For example, internalized protease-activated receptors (PAR1) are specifically sorted to lysosomes and do not recycle (18).

Similar to the ß2AR, the angiotensin II type 1A receptor (AT1AR) is rapidly phosphorylated and internalized after its activation with agonist (19, 20, 21). However, the agonist-stimulated internalization of the AT1AR appears to be more complex than the internalization of the ß2AR. While the predominant mechanism(s) governing AT1AR remain unclear, AT1AR internalization in HEK 293 and COS7 cells is not sensitive to inhibition by dominant-negative proteins of ß-arrestin and dynamin, which interfere with clathrin-coated vesicle-mediated internalization (9). Furthermore, the AT1AR colocalizes with ß-arrestin in endocytic vesicles, a property that is not shared by the ß2AR (22). Nonetheless, there are conflicting reports in the literature regarding whether or not AT1AR internalization can be blocked using dominant-negative ß-arrestin and dynamin mutants, suggesting that ß-arrestins and clathrin may play a more substantial role in regulating AT1AR internalization than originally envisaged (9, 20, 23).

Recently, using ß2AR/AT1AR chimeric receptors and ß-arrestin2 fused to green fluorescent protein (GFP), we demonstrated that the carboxyl-terminal tails of these receptors determined whether or not ß-arrestin redistributed to large core intracellular vesicular structures in response to receptor activation (22). However, it is unknown whether ß-arrestin is physically associated with the AT1AR in endocytic vesicles. Therefore, we tested whether the carboxyl-terminal tails of the ß2AR and AT1AR, in part via differences in their ability to form stable complexes with ß-arrestin, contribute to differences in endocytosis, dephosphorylation, and plasma membrane-recycling patterns observed for the ß2AR and AT1AR.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Coimmunoprecipitation of ß2AR-ATCT/ß-Arrestin2 Complexes
Previously, we demonstrated that agonist activation of the AT1AR, but not the ß2AR, resulted in the redistribution of ß-arrestin2-GFP to large core intracellular vesicular structures (22). Furthermore, we demonstrated that, by replacing the carboxyl-terminal tail of the ß2AR with the AT1AR tail, the ß2AR gained the capacity to stimulate the redistribution of ß-arrestin2-GFP fluorescence to endocytic vesicles (22). While these observations are suggestive of the formation of AT1AR/ß-arrestin2-GFP complexes in endosomes, the evidence supporting this association is circumstantial and does not demonstrate a physical association between ß-arrestin and the receptor tail in subcellular membrane locations. Therefore, to determine whether ß-arrestin2-GFP is physically associated with the AT1AR carboxyl-terminal tail in endocytic vesicles, we attempted to coimmunoprecipitate wild-type ß2AR and ß2AR-ATCT chimera with ß-arrestin2-GFP from both plasma membrane and light vesicular membrane fractions. Isoproterenol stimulation of HEK 293 cells expressing ß2AR resulted in a 2.6 ± 1.1-fold increase in ß2AR co-immunoprecipitated with ß-arrestin2-GFP from the plasma membrane fraction vs. cells that were not exposed to agonist (Fig. 1AGo). Furthermore, the amount of ß2AR-ATCT and ß2AR coimmunoprecipitated with ß-arrestin2-GFP from the plasma membrane fraction after isoproterenol treatment was not apparently different, 2.2 ± 0.6- vs. 2.6 ± 1.1-fold, respectively (Fig. 1AGo). However, in the absence of isoproterenol, the amount of ß2AR-ATCT coimmunoprecipitated with ß-arrestin2-GFP from the plasma membrane fraction was 1.7 ± 0.5-fold greater than that coimmunoprecipitated with the wild-type receptor (Fig. 1AGo). ß2AR coimmunoprecipitated with ß-arrestin2-GFP from the light vesicular membrane fraction was not increased by isoproterenol stimulation (Fig. 1BGo). However, agonist treatment of ß2AR-ATCT-expressing cells resulted in a 2.7 ± 1-fold increase in ß2AR-ATCT coimmunoprecipitated with ß-arrestin2-GFP from the vesicular fraction (Fig. 1BGo). These experiments demonstrate that agonist promotes the association of ß-arrestin2-GFP with both the ß2AR and ß2AR-ATCT at the plasma membrane, but that the localization of ß-arrestin2 to endocytic vesicles requires that it physically associate with the AT1AR carboxyl-terminal tail of the ß2AR-ATCT chimera.



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Figure 1. Agonist-Dependent Coimmunoprecipitation of the ß2AR and ß2AR-ATCT with ß-Arrestin2-GFP

A, The relative amount of ß2AR and ß2AR-ATCT immunoprecipitated with ß-arrestin2-GFP from the plasma membrane fraction of HEK 293 cells incubated in the absence and presence of 10 µM isoproterenol for 45 min at 37 C. B, The relative amount of of ß2AR and ß2AR-ATCT immunoprecipitated with ß-arrestin2-GFP from the light vesicular membrane fraction of HEK 293 cells incubated in the absence and presence of 10 µM isoproterenol for 45 min at 37 C. The preparation of subcellular membrane fractions and 125I-pindolol binding to receptors coimmunoprecipitated with ß-arrestin2-GFP using a rabbit polyclonal anti-GFP antibody were performed as described in Materials and Methods. The data represent the fold increase in receptor immunoprecipitated when compared with ß2AR immunoprecipitated with ß-arrestin2-GFP in the absence of agonist stimulation in each individual experiment. The data represent the mean ± SD of three independent experiments. *, P < 0.05 vs. ß2AR without agonist.

 
Recently, Oakley et al. (24) demonstrated that the redistribution of ß-arrestin to vesicles with the vasopressin receptor required GRK-mediated phosphorylation of a cluster of serine residues within the carboxyl-terminal tail of the receptor. Therefore, we prepared two AT1AR truncation mutants to determine whether the deletion of two clusters of putative sites for GRK-mediated phosphorylation in the carboxyl-terminal tail of the AT1AR (amino acid residues 326–338 and 342–348) contribute to the localization of ß-arrestin to endocytic vesicles. In response to agonist stimulation, ß-arrestin2-GFP becomes completely localized to large core vesicles in cells expressing wild-type AT1AR (Fig. 2AGo). In cells expressing an AT1AR mutant ({Delta}339) lacking the final 20 amino acid residues of the carboxyl-terminal tail, ß-arrestin2-GFP demonstrates a punctate distribution at the plasma membrane surface and is localized to small endocytic vesicles after agonist treatment (Fig. 2BGo). ß-Arrestin2-GFP distribution after agonist activation of an AT1AR mutant ({Delta}319) lacking the final 40 amino acid residues of the carboxylterminal tail is limited to a punctate distribution at the plasma membrane (Fig. 2CGo). Each of the truncated receptors internalized in response to agonist stimulation (data not shown). Thus, while ß-arrestin2 translocation and redistribution to coated pits in response to AT1AR activation does not require the association of ß-arrestin with the AT1AR carboxyl-terminal tail, the tail is required for ß-arrestin2-GFP redistribution to endocytic vesicles.



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Figure 2. Effect of Truncating the AT1AR Carboxyl-Terminal Tail on Agonist-Stimulated ß-Arrestin2-GFP Redistribution

ß-Arrestin2-GFP distribution in AT1AR (panel A), AT1AR-{Delta}339 (panel B), and AT1AR-{Delta}319 (panel C) expressing HEK 293 cells after 45 min treatment with 100 nM angiotensin II. HEK 293 cells were transfected transiently with plasmid cDNAs encoding HA epitope-tagged receptors (10 µg) and pEGFP-N1 ß-arrestin2-GFP (5 µg). Data shown are representative of three experiments. Bar = 10 µm.

 
Clathrin Dependence of AT1AR/ß-Arrestin2 Complex Formation
Since AT1AR endocytosis is mediated by both dynamin-dependent and -independent mechanisms and is phosphorylation dependent (9, 20, 23), we tested whether the formation of AT1AR/ß-arrestin2-GFP complexes in endosomes is a dynamin-dependent process. In the absence of agonist, ß-arrestin2-GFP is diffusely localized throughout the cytoplasm of cells expressing the wild-type AT1AR (Fig. 3AGo), and, upon agonist activation, ß-arrestin2-GFP becomes localized to large core intracellular vesicles (Fig. 3BGo). Unexpectedly, in cells expressing both the AT1AR and dynamin I-K44A, ß-arrestin2-GFP fluorescence was localized to the plasma membrane in a punctuated distribution even in the absence of agonist stimulation (Fig. 3CGo). Moreover, in the presence of dynamin I-K44A, no redistribution of ß-arrestin2-GFP to the endosomal compartment was observed after a 45-min exposure to 100 nM angiotensin II (Fig. 3DGo). In contrast, in cells that expressed the AT1AR-ß2CT chimera, dynamin I-K44A overexpression did not alter the cytosolic distribution of ß-arrestin2-GFP (Fig. 3Go, panel E vs. panel G) and did not prevent AT1AR-ß2CT-mediated ß-arrestin2-GFP translocation to the cell surface in response to agonist stimulation (Fig. 3Go, F and H). Consistent with the idea that the formation of ß-arrestin/AT1AR complexes is clathrin dependent, clathrin immunofluorescence colocalized with ß-arrestin2-GFP in large core vesicles in AT1AR- but not ß2AR-expressing cells (Fig. 4Go). Taken together, these observations indicate that the endocytosis of AT1AR/ß-arrestin2 complexes is mediated by clathrin-coated vesicles and not by the putative dynamin-independent endocytic pathway previously described for the AT1AR (9). Furthermore, the localization of ß-arrestin2-GFP to the plasma membrane in the presence of dynamin I-K44A, in AT1AR- but not AT1AR-ß2CT-expressing cells, suggests that ß-arrestins can associate with the AT1AR carboxyl-terminal tail in the absence of agonist activation.



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Figure 3. Effect of Dynamin I-K44A Overexpression on ß-Arrestin2-GFP Translocation and Intracellular Redistribution in Response to AT1AR and AT1AR-ß2CT Activation

ß-Arrestin2-GFP distribution in AT1AR- and AT1AR-ß2CT-expressing HEK 293 cells in the absence of dynamin I-K44A before (A and E) or after (B and F) 30 min exposure of the same cells to 100 nM angiotensin II. Effect of dynamin I-K44A overexpression on ß-arrestin2-GFP distribution in AT1AR- and AT1AR-ß2CT-expressing HEK 293 cells before (C and G) or after (D and H) 30 min exposure of the same cells to 100 nM angiotensin II. HEK 293 cells were transfected transiently with plasmid cDNAs encoding HA epitope-tagged receptors (5–10 µg) and pEGFP-N1 ß-arrestin2-GFP (5 µg) with or without plasmid cDNAs encoding dynamin I-K44A (5 µg). Data shown are representative of four experiments. Bar represents 10 µm.

 


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Figure 4. Agonist-Stimulated Colocalization of ß-Arrestin2-GFP and Clathrin in HEK 293 Cells

Confocal visualization of the distribution and colocalization (overlay) of ß-arrestin2-GFP (green) and X-22 monoclonal antibody-labeled clathrin (red) in HEK 293 cells expressing ß2AR (panel A) and AT1AR (panel B). Shown are representative images of cells fixed in 3.6% paraformaldehyde after a 45-min stimulation with either 10 µM isoproterenol or 100 nM angiotensin II. Cells were transfected with 10 µg of plasmids encoding either ß2AR or AT1AR along with 5 µg of ß-arrestin2-GFP. Data shown are representative of four experiments. Bar represents 10 µm.

 
To further establish that the association of ß-arrestin with the AT1AR carboxyl-terminal tail mediates the agonist-independent membrane localization of ßarrestin2-GFP, we examined whether dynamin I-K44A overexpression altered ß-arrestin2-GFP distribution in cells expressing either the wild-type ß2AR or ß2AR-ATCT chimera. In cells expressing the wild-type ß2AR with or without dynamin I-K44A, ß-arrestin2-GFP fluorescence is diffusely distributed throughout the cytoplasm in the absence of agonist and translocates to the plasma membrane in response to agonist activation of the receptor (Fig. 5Go, A–D). However, in cells expressing the ß2AR-ATCT chimera, ß-arrestin2-GFP is localized at the plasma membrane surface (>75% of cells) even in the absence of dynamin I-K44A expression (Fig. 5Go, E and G). This observation is consistent with data presented in Fig. 1Go where 1.7 ± 0.5-fold more ß2AR-ATCT than ß2AR is coimmunoprecipitated with ß-arrestin2-GFP in the absence of agonist stimulation. However, the redistribution of ß-arrestin2-GFP to large core intracellular vesicular structures requires agonist activation of the receptor (Fig. 5Go, panel E vs. panel F). Similar to what is observed for the AT1AR, dynamin I-K44A blocked the clathrin-coated vesicle-mediated endocytosis of ß2AR-ATCT/ß-arrestin2-GFP complexes (Fig. 5HGo).



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Figure 5. Effect of Dynamin I-K44A Overexpression on ß-Arrestin2-GFP Translocation and Intracellular Trafficking in Response to either ß2AR or ß2AR-ATCT Activation

ß-Arrestin2-GFP distribution in both ß2AR- and ß2AR-ATCT-expressing HEK 293 cells in the absence of dynamin I-K44A before (A and E) or after (B and F) 30 min exposure of the same cells to 10 µM isoproterenol. Effect of dynamin1-K44A overexpression on ß-arrestin2-GFP distribution in both ß2AR- and ß2AR-ATCT-expressing HEK 293 cells before (C and G) or after (D and H) 30 min exposure of the same cells to 10 µM isoproterenol. HEK 293 cells were transfected transiently with plasmid cDNAs encoding pEGFP-N1 ß-arrestin2-GFP (5 µg) and either HA epitope-tagged pcDNA1-Amp ß2AR (10 µg) or FLAG epitope-tagged pcDNA1-Amp ß2AR-ATCT (10 µg) with or without plasmid cDNAs encoding dynamin I-K44A (5 µg). Inset shows the redistribution of ß-arrestin2-GFP to large-core intracellular vesicles. Data shown are representative of three independent experiments. Bar represents 10 µm.

 
Agonist-Independent Receptor Internalization
The localization of ß-arrestin2-GFP fluorescence at the plasma membrane surface in the absence of agonist suggested that the association of ß-arrestin with the AT1AR carboxyl terminus might promote agonist-independent ß2AR-ATCT and AT1AR internalization. Moreover, agonist-independent internalization of the AT1AR has previously been reported (25). Therefore, we tested for the loss of cell surface ß2AR-ATCT and AT1AR in the absence of agonist stimulation. Epitope-tagged receptors were labeled with primary antibody at 4 C for 45 min, and the loss of cell surface secondary antibody labeling was determined after the warming of cells to 37 C in the absence of agonist. Significant loss of cell surface ß2AR-ATCT was observed (VMAX = 40 ± 6) in the absence of agonist stimulation and was not increased by the overexpression of ßarrestin2 (Fig. 6AGo). In contrast, agonist-independent internalization of the wild-type AT1AR was not observed in the absence of overexpressed ß-arrestin2 (VMAX = 2 ± 1%) but was observed in the presence (VMAX = 31 ± 9%) of overexpressed ß-arrestin2 (Fig. 6AGo). Moreover, no agonist-independent AT1AR-ß2CT internalization was observed in the presence or absence of overexpressed ß-arrestin2 (data not shown). To examine whether endogenous ß-arrestins contributed directly to the agonist-independent loss of cell surface ß2AR-ATCT, we tested the ability of a dominant-negative ß-arrestin mutant, ß-arrestin1 (185–418),to inhibit agonist-independent ß2AR-ATCT endocytosis. When tested, ß-arrestin1 (185–418) significantly retarded the agonist-independent loss of cell surface ß2AR-ATCT (VMAX = 34 ± 4% vs. 56 ± 5%, P < 0.05) (Fig. 6BGo). The agonist-independent loss of cell surface ß2AR-ATCT was unaffected by the treatment of cells with propranolol (10 µM) (Fig. 6BGo). These experiments demonstrate that ß-arrestin-dependent receptor internalization can occur in the absence of receptor activation by agonist.



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Figure 6. Agonist-Independent Endocytosis of the AT1AR and ß2AR-ATCT in the Presence and Absence of Overexpressed Wild-Type and Dominant-Negative ß-Arrestins

A, Time course for the agonist-independent internalization of the AT1AR (circles) and ß2AR-ATCT (squares) in the absence (open symbols) and presence of overexpressed ßarrestin2 (filled symbols). B, Time course for the agonist-independent endocytosis of the ß2AR-ATCT in the absence or presence of either ß-arrestin1 (185–418), or 10 µM propranolol. Cell surface receptors were labeled with primary antibody on ice as described in the Materials and Methods and then warmed to 37 C for the times indicated. Agonist-independent internalization is defined as a loss of antibody-labeled cell surface receptor sites in the absence of agonist stimulation. HEK 293 cells were transfected transiently with plasmid cDNA containing 12CA5-AT1AR (1–2 µg) and FLAG-ß2AR-ATCT (1–2 µg) and either empty vector (10 µg), wild-type ß-arrestin2 (10 µg), or ß-arrestin1 mutants (10 µg). The data represent the mean ± SE of three independent experiments normalized to the loss of ß2AR cell surface receptor in the absence of agonist stimulation.

 
Wild-Type and Chimeric Receptor Recycling
While it is well documented that internalized ß2AR are recycled back to the cell surface (13, 14, 15), it is less clear whether this occurs for the AT1AR. Therefore, using both wild-type and chimeric ß2AR and AT1AR, we examined whether differences exist in the relative ability of these receptors to recycle and whether this might correlate with their observed capacity to form complexes with ß-arrestin complexes in endosomes. When tested, we found that, unlike the ß2AR, AT1AR were not efficiently recycled back to the cell surface after the washout of agonist even in the absence of ß-arrestin overexpression (Fig. 7Go, A and B). However, when the AT1AR carboxyl-terminal tail was replaced with the ß2AR tail, plasma recycling of the AT1AR was observed (Fig. 7BGo), whereas the replacement of the ß2AR carboxyl-terminal tail with the AT1AR tail prevented plasma membrane recycling of the ß2AR (Fig. 7AGo). These results suggest that the plasma membrane recycling of ß2AR and AT1AR is negatively correlated with the formation of receptor/ß-arrestin complexes in endosomes.



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Figure 7. Role of the ß2AR and AT1AR Carboxyl-Terminal Tails in Receptor Recycling

Time courses for the recycling of the wild-type ß2AR and the ß2AR-ATCT chimera (panel A) and the wild-type AT1AR and the AT1AR-ß2CT chimera (panel B) to the cell surface after agonist stimulation for 30 min with 10 µM isoproterenol and 100 nM angiotensin II, respectively. Cells were washed twice on ice and then incubated at 37 C in the absence of agonist for the times indicated in the figure. Receptor recycling was measured as a time-dependent return of cell surface immunofluorescence after agonist removal. HEK 293 cells were transfected transiently with plasmid cDNA containing the 12CA5-ß2AR (250 ng), 12CA5-AT1AR (1–2 µg), FLAG-ß2AR-ATCT (1–2 µg), and 12CA5 AT1AR-ß2CT (1–2 µg) along with either empty vector (1 µg) or ß-arrestin2 (1 µg) in the case of the ß2AR-ATCT. The extent of internalization after 30 min exposure to agonist for each receptor in these experiments was as follows: ß2AR = 27 ± 2%, ß2AR-ATCT = 28 ± 2%, AT1AR = 72 ± 4%, and AT1AR-ß2CT = 66 ± 1% of cell surface receptors. The data represent the mean ± SE of three experiments.

 
Wild-Type and Chimeric Receptor Dephosphorylation
In the case of the ß2AR, internalization is required for receptor dephosphorylation and resensitization (14, 17). However, it is unknown whether internalized AT1AR are dephosphorylated. Therefore, we examined the whole cell phosphorylation and dephosphorylation of the ß2AR and AT1AR in the absence of ß-arrestin overexpression. Both receptors were effectively phosphorylated in response to agonist stimulation (Fig. 8Go, A and B). When agonist was removed and cells were allowed to recover in agonist-free media for 20 min, marked dephosphorylation was observed for the wild-type ß2AR, but not the wild-type AT1AR (Fig. 8Go, A and B). To test whether ß-arrestin dissociation from the ß2AR was required for dephosphorylation, similar to that observed previously for rhodopsin (26), we tested the dephosphorylation of the ß2AR-ATCT chimera. We find that the ß2AR-ATCT chimera, which exhibits the capacity to form a complex with ß-arrestin in endosomes, did not become dephosphorylated when allowed to recover in agonist-free medium (Fig. 8AGo). However, the AT1AR-ß2CT chimera, which does not exhibit the capacity to internalize with ß-arrestin bound, also did not become dephosphorylated (Fig. 8BGo). The lack of AT1AR-ß2CT dephosphorylation indicates that the lack of AT1AR dephosphorylation may be independent of ß-arrestin complex formation. This suggests that receptor dephosphorylation is not common to all GPCRs.



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Figure 8. Whole-Cell Phosphorylation and Dephosphorylation of Wild-Type and Chimeric ß2AR and AT1AR

HEK 293 transfected to overexpress ß2AR and ß2AR-ATCT (panel A) and AT1AR and AT1AR-ß2CT (panel B) were labeled with 100 µCi/ml [32P]orthophosphate for 1 h in phosphate-free medium. Cells were pretreated for 10 min in serum-free medium at 37 C in the absence (N, naive) or presence (D, desensitized; R, resensitized) of either 10 µM isoproterenol or 100 nM angiotensin II, washed three times, and either allowed to resensitize (R) for 20 min at 37 C or kept on ice (N, D). Representative autoradiographs and phosphorimager analysis of the mean ± SEM of three independent experiments demonstrating the whole-cell phosphorylation of the ß2AR and ß2AR-ATCT (panel A) and AT1AR and AT1AR-ß2CT (panel B) are shown. Each lane was loaded with equivalent amounts of receptor protein as described in Materials and Methods. The phosphorimager data are normalized to receptor phosphorylation in the absence of agonist. *, P < 0.05 vs. desensitized wild-type receptor phosphorylation.

 
AT1AR Desensitization and Resensitization
AT1AR-mediated GFP-protein kinase C (PKC)ßII translocation was previously demonstrated to serve as an effective measure of homologous AT1AR desensitization (27). This assay was used to determine receptor desensitization/resensitization in the present experiments for two reasons: 1) the sensitivity of the assay and 2) the determination of receptor desensitization/resensitization is not confounded by the accumulation of intracellular second messengers in response to the desensitizing stimulus. In cells expressing either the wild-type or chimeric AT1AR, angiotensin II stimulated the plasma membrane translocation of GFP-PKCßII (Fig. 9AGo). In both cases, the GFP-PKCßII translocation response desensitized rapidly (Fig. 9AGo). However, while GFP-PKCßII translocation was reestablished in cells that were allowed to recover for 20 min in agonist-free media (Fig. 9BGo), the magnitude of GFP-PKCßII plasma membrane translocation in response to agonist remained attenuated for both the AT1AR and the AT1AR-ß2CT chimera when compared with untreated control cells (Fig. 9CGo).



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Figure 9. Activation, Desensitization, and Resensitization of AT1AR- and AT1AR-ß2CT-Mediated PKC-GFP Translocation

A, Time course for the activation and desensitization of AT1AR and AT1AR-ß2CT-mediated GFP-PKCßII membrane translocation in responses to 100 nM angiotensin II. Data shown are representative of seven independent experiments. B, Resensitization of AT1AR- and AT1AR-ß2CT-mediated PKC-GFP translocation responses to 100 nM angiotensin II after the washout of agonist and a 30-min recovery of cells in agonist-free media. The data are representative of five to six different experiments. C, Quantification of the relative magnitude of AT1AR- and AT1AR-ß2CT-stimulated GFP-PKCßII translocation responses in control cells and cells allowed to resensitize 30 min in the absence of agonist after the preexposure of cells to 100 nM angiotensin II. The relative GFP-PKCßII fluorescence intensity and duration of membrane translocation were measure using the LSM-510 image analysis software and the normalized data presented as comparison with data obtained from control AT1AR-expressing cells. The data for each curve represent the mean ± SEM of the data obtained from 13 different AT1AR-expressing cells from seven distinct experiments and 17–20 AT1AR-ß2CT-expressing cells from five to six independent experiments. *, P < 0.05 vs. control. Bar represents 10 µm.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Our findings provide direct biochemical and cell biological evidence that the redistribution of ß-arrestin2 to large-core intracellular vesicular structures in response to AT1AR activation involves the formation of a stable complex between the carboxyl-terminal tail of the receptor and ß-arrestin2. In contrast, the ß2AR carboxyl-terminal tail does not support the formation of intracellular receptor/ß-arrestin complexes. We find that the stable association of ß-arrestin with the AT1AR tail not only mediates the agonist-stimulated internalization of the AT1AR via clathrin-coated pits but also has the potential to allow the agonist-independent internalization of the AT1AR via clathrin-coated vesicles. Furthermore, while internalized ß2AR is recycled back to the cell surface, internalized AT1AR are retained within the cell and do not recycle. However, the relative ability of these receptors to recycle appears to be regulated by their carboxyl-terminal tails and is correlated with the ability of the receptor tails to form stable complexes with ß-arrestin. In the case of the ß2AR, ß-arrestin dissociation was required for receptor dephosphorylation in the endosomal compartment but, surprisingly, dephosphorylation of the AT1AR did not occur even under conditions where no ß-arrestin association with the receptor in endosomes was expected to occur. This observation suggests that receptor dephosphorylation may not represent a property common to all GPCRs. Taken together, these data support the hypothesis that receptor carboxyl-terminal tails not only regulate the desensitization of GPCRs, but also play an essential role in regulating differences in receptor trafficking and resensitization patterns. In part, this may be the consequence of their relative capacity to form stable complexes with ßarrestin proteins.

To our knowledge, the present experiments provide the first evidence implicating ß-arrestins in promoting agonist-independent GPCR endocytosis. Although agonist-stimulated GPCR internalization is now a well established phenomenon (1, 2, 3), the concept that GPCRs internalize in an agonist-independent manner is less well accepted. Agonist-independent receptor internalization has now been reported for a growing number of GPCRs, including the AT1AR (25, 28, 29). While we observed agonist-independent AT1AR internalization only after ß-arrestin overexpression, Hein et al. (25) documented agonist-independent AT1AR internalization in HEK 293 cells in the absence of ß-arrestin overexpression. This discrepancy suggests that there exists heterogeneity in ß-arrestin expression levels between different HEK 293 cell cultures. Nonetheless, it is likely that the agonist-independent internalization of some GPCRs will be found to be ß-arrestin independent.

While our data show that ß-arrestins can associate with receptors in the absence of agonist occupancy, agonist stimulation clearly stabilizes the complex formed between the receptor and ß-arrestin, since ß-arrestin redistribution to endosomes was observed only in the presence of agonist (Fig. 4Go, compare panels E and F). The stabilization of the receptor/ß-arrestin complex by agonist is likely the consequence of both receptor isomerization to a "high-affinity" ligand binding state and GRK-mediated phosphorylation of the receptor C-terminal tail. This is consistent with a recent report demonstrating that vasopressin V2 receptor-stimulated redistribution of ß-arrestin to endosomes involved GRK2 phosphorylation of a triplet motif of serine residues in the carboxyl-terminal tail of the receptor that presumably stabilized interactions of ß-arrestin with the receptor (24). In the case of PAR1, the mutation of putative receptor phosphorylation sites in the carboxyl-terminal tail of PAR1 prevented agonist-stimulated PAR1 internalization, whereas agonist-independent PAR1 internalization was unaffected by these mutations (28). Our data suggest that, under the appropriate conditions, the agonist-independent association of ß-arrestin with both the ß2AR-ATCT mutant and AT1AR is facilitated by a receptor conformation promoted by the AT1AR carboxyl-terminal tail. The agonist-independent association of ßarrestin with these receptors is not supported by the ß2AR carboxyl-terminal tail, which may negatively regulate the association of ß-arrestin in the absence of agonist stimulation. The higher-affinity agonist-independent interactions with the ß2AR-ATCT mutant may reflect receptor-specific differences in the contribution of other intracellular domains to ß-arrestin interactions. Nonetheless, for the AT1AR, ß-arrestin regulates both agonist-stimulated and agonist-independent internalization. It is possible that the role of GRK phosphorylation in the formation of stable receptor/ßarrestin complexes may represent the molecular event distinguishing agonist-stimulated PAR1 internalization to lysosomes from the agonist-independent PAR1 trafficking (28).

In addition to the AT1AR, receptor-mediated redistribution of ß-arrestin to intracellular endocytic structures has recently been reported to occur in response to neurotensin, vasopressin V2, neurokinin 1, TRH, and protease receptor activation (22, 24, 30, 31, 32). However, while each of these receptors is presumed to be associated with ß-arrestin in endocytic vesicles, there is no evidence supporting a direct interaction between any of these receptors and ß-arrestin in endosomes. In the present study, we show that ß-arrestin2 is physically associated with the AT1AR carboxyl-terminal tail and that the AT1AR tail, through its association with ß-arrestin, potentially prevents receptor recycling. Nonetheless, it remains unclear whether ß-arrestin acts as a retention signal preventing receptor recycling. However, except for the neurokinin 1 receptor (15, 32), poor recycling to the plasma membrane is a common property shared by receptors that stimulate ß-arrestin redistribution to intracellular vesicles (24, 30, 31, 33). The fact that the neurokinin 1 receptor recycles back to the cell surface suggests that either this receptor is internalized to a distinct endosomal compartment or that the neurokinin 1 receptor does not physically associate with ß-arrestin in endocytic vesicles. Moreover, it is also likely that the specific association of distinct GPCRs with additional components of the endosomal compartment may also contribute to differences in both the intracellular trafficking patterns and extent of plasma membrane recycling for each of these GPCRs. The finding that, unlike observed for the vasopressin V2 receptor (24), the ß2AR tail does not completely rescue AT1AR recycling may suggest that other receptor domains contribute to the intracellular retention of the AT1AR. The answers to these questions remain to be determined.

In the present study, we provide evidence that in response to agonist, ß-arrestin is physically associated rather than colocalized with the AT1AR in endocytic vesicles. First, we show that the ß2AR bearing the AT1AR tail can be coimmunoprecipitated with ß-arrestin from the light vesicular membrane pool. Second, the loss of all potential sites for GRK-mediated AT1AR phosphorylation does not prevent ß-arrestin association with AT1AR but completely prevents the localization of ß-arrestin to endocytic vesicles. Third, the coexpression of the dynamin I-K44A inhibitor of clathrin-mediated endocytosis prevented the redistribution of ß-arrestin to endocytic vesicles. Finally, by switching the carboxyl-terminal tails between the AT1AR and the ß2AR, we reversed the ability of these receptors to internalize with ß-arrestin without preventing their ability to stimulate ß-arrestin translocation. These findings are consistent with other data showing that phosphorylation is required for the agonist-dependent redistribution of ß-arrestin with both the vasopressin V2 receptor and AT1AR to endocytic vesicles (Ref. 24 and R. H. Oakley and M. G. Caron, personal communication). Taken together, these observations suggest that the signal for stable GPCR/ß-arrestin association involves phosphorylation and that ß-arrestins remain associated with the receptors as they traffic through the endosomal compartment. Future experiments, using bioluminescent resonance energy transfer (BRET) will be required to determine whether the association of ß-arrestin with receptors in the endosomal compartment is either static or dynamically regulated (34).

Recently, Oakley et al. (35) analyzed arrestin translocation in response to the activation of multiple different GPCRs. In doing so, they identified two distinct classes of GPCRs: class A GPCRs (e.g. ß2AR), receptors that interact with ß-arrestin2 with greater affinity than ß-arrestin1 and do not interact with visual arrestin, and class B GPCRs (e.g. AT1AR), receptors that bind equally well to both ß-arrestin isoforms and also associated with visual arrestins. Furthermore, using ß2AR/vasopressin V2 receptor chimeras, Oakley et al. (35) demonstrated that the carboxyl-terminal tails of the receptors determined the affinity of receptors for ß-arrestins and visual arrestins. Interestingly, each of the class B receptors promotes the redistribution of ß-arrestin to endocytic vesicles (24). In the case of at least two class B GPCRs, the vasopressin V2 receptor and AT1AR, GRK-mediated phosphorylation is required for ß-arrestin redistribution to endocytic vesicles. Our observation that deleting the tail of the AT1AR to remove putative sites for GRK-mediated phosphorylation (residues 326–338) does not prevent ß-arrestin translocation in response to AT1AR activation suggests that phosphorylation stabilizes the complex between the receptor and ß-arrestin. Thus, multiple domains contribute to the association of ß-arrestins with GPCRs, but the tail appears to regulate the affinity of these interactions. Consequently, the observed differences in ß-arrestin affinity that is observed for class A vs. class B receptors may be determined by differences in the patterns and/or extent of GRK-mediated phosphorylation.

An important facet of ß2AR internalization is to promote receptor dephosphorylation and resensitization. We find that the internalization of ß-arrestin with the ß2AR-ATCT prevented the dephosphorylation of the ß2AR chimera, indicating that receptor dephosphorylation in the endosomal compartment appears to require the dissociation of the receptor/ß-arrestin complex (Fig. 8Go). However, the AT1AR-ß2CT chimera, which does not internalize with ß-arrestin bound, was not dephosphorylated. It is proposed that acidification of receptors in endosomes induces a conformational change in the receptor that supports the association of a GPCR-specific phosphatase (16, 36). This suggests that, in addition to the dissociation of the receptor/ß-arrestin complex, receptors must exhibit the capacity to serve as phosphatase substrates. It would appear that either the AT1AR lacks domains required for the binding of phosphatases or acidification in endocytic vesicles does not induce receptor conformational changes in AT1AR structure required for phosphatase association. Alternatively, the lack of AT1AR-ß2CT dephosphorylation might be the consequence of a fortuitous interaction between intracellular domains. However, all other receptor properties can be exchanged normally between the receptors.

The data presented in this paper suggest the following model for the intracellular trafficking and resensitization of the ß2AR and AT1AR (Fig. 10Go). In the absence of inhibitors, such as dynamin I-K44A, agonist stimulation promotes the desensitization and internalization of both the ß2AR and AT1AR by a common mechanism, GRK phosphorylation and ß-arrestin binding. The AT1AR internalizes with ß-arrestin bound, whereas ß-arrestin dissociates from the ß2AR shortly after the formation of endocytic vesicles. In the case of the ß2AR, acidification of the receptor in endocytic vesicles enhances receptor dephosphorylation by a membrane-bound receptor-specific phosphatase (16, 36), after which the receptor is rapidly recycled back to the cell surface. In the case of the AT1AR, internalized receptor is not dephosphorylated in endocytic vesicles due to both the continued association of the receptor with ß-arrestin and the inability of the receptor to serve as a phosphatase substrate. The resensitization of AT1AR-mediated responses is mediated by either the delayed recycling of internalized receptors back to the cell surface, the surface reexpression of receptors that were internalized in the absence of agonist stimulation, or the mobilization of newly synthesized receptors to the cell surface (25, 28).



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Figure 10. Proposed Model for Difference in the Intracellular Trafficking and Surface Reexpression of ß2AR and AT1AR

Agonist activation of ß2AR and AT1AR in HEK 293 cells leads to their desensitization and internalization by a common mechanism involving GRK phosphorylation and ßarrestin binding. Both receptors are internalized by clathrin-coated pits, and internalized ß2AR is dephosphorylated and rapidly recycled back to the cell surface, whereas the AT1AR internalizes with ßarrestin and is neither dephosphorylated nor recycled. AT1AR resensitization is either mediated by the slow recycling of receptor back to the cell surface, de novo receptor synthesis, or the recruitment of a reserve pool of intracellular receptors. ßArr, ßarrestin; G, G protein; GRK, G protein-coupled receptor kinase; and GRP, G protein-coupled receptor phosphatase; A, agonist.

 
In summary, while many of the molecular mechanisms first described using the ß2AR as a model apply to other GPCRs (1, 2, 3), the recent characterization of a diverse variety of GPCRs has revealed clear differences in the intracellular trafficking agendas and resensitization patterns of many GPCRs (9, 18, 22, 24, 30, 31, 32, 33, 35, 37). In the present study, we have demonstrated that the carboxyl-terminal receptor domains not only contribute to the endocytosis of the ß2AR and AT1AR, but that the stable association of ß-arrestin with the AT1AR carboxyl-terminal tail contributes to observed differences in the intracellular trafficking, surface reexpression, and resensitization patterns of these receptors. Therefore, while GPCR activation, desensitization, and internalization may involve common mechanisms, observed differences in receptor responsiveness may involve differences in the intracellular trafficking patterns of GPCRs that result from differences in their capacity to form stable complexes with their regulatory proteins.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Materials
Human embryonic kidney cells (HEK 293) were provided by the American Type Culture Collection (ATCC, Manassas, VA). Tissue culture media and FBS were obtained from Life Technologies, Inc. (Gaithersburg, MD). Isoproterenol was purchased from Research Biochemicals International (Natick, MA). Mouse anti-HA 12CA5 and anti-Flag monoclonal antibodies were obtained from Roche Molecular Biochemicals (Indianapolis, IN) and Research Diagnostic, Inc. (Flanders, NJ), respectively. 125I-Pindolol was purchased from NEN Life Science Products (Boston, MA). fluorescein isothiocyanate (FITC)-conjugated goat antimouse secondary antibody and angiotensin II and all other biochemical reagents were purchased from Sigma.

DNA Construction
All recombinant cDNA procedures were carried out following standard protocols. The ßarrestin1 mutant, 185–418, was constructed by PCR. 5'-Oligonucleotide primers introduced an amino-terminal EcoRI restriction site, minimal Kozak sequence, and initiation ATG at the appropriate site of ß-arrestin1, and 3'-oligonucleotide primers introduced a carboxyl-terminal XhoI restriction site, stop codon, and Flag-epitope tag sequence (DYKDDDDK) at the C terminus of ß-arrestin1. The AT1AR carboxyl-terminal tail truncation mutants were also constructed by PCR. 3'-Oligonucleotide primers were used to introduce a stop codon after tyrosine residues 319 and 339 followed by a XbaI restriction site. The 5'-oligonucleotide was identical to the AT1AR sequence upstream of an unique EcoRI site. The resulting PCR products were digested with EcoRI and XbaI and subcloned into the pcDNA3.1 AT1AR plasmid construct digested with the same enzymes. The sequence integrity of the ß-arrestin and AT1AR mutants was confirmed by DNA sequencing. All other cDNA constructs used have been reported previously (8, 22, 27).

Cell Culture and Transfection
HEK 293 cells were grown in Eagle’s minimal essential medium with Earle’s salt (MEM) supplemented with 10% (vol/vol) heat-inactivated FBS and gentamicin (100 µg/ml). The cells were seeded at a density of 2.5 x 106/100-mm dish and transiently transfected with the cDNAs described in the figure legends by a modified calcium phosphate method. After transfection (~18 h) the cells were reseeded into 35-mm glass-bottomed culture dishes (MarTek) for confocal studies or six-well Falcon dishes.

Receptor Expression
Receptor expression was 1,000–3,000 fmol/mg whole-cell protein for confocal studies and 1,000–1,500 fmol/mg whole-cell protein for internalization and phosphorylation studies. AT1AR expression was matched to ß2AR expression by flow cytometry, and ß2AR expression was measured by saturating 125I-pindolol binding (8, 9, 38).

Subcellular Fractionation
HEK293 cells transfected with either ß2AR or ß2AR-ATCT were plated in 100-mm dishes, treated as described in the figure legends, 1.8 x 107 cells were scraped in 6 ml of 10 mM Tris, pH 7.4, with 1 mM EDTA, dounce homogenized on ice (20 strokes), and spun at 500 x g for 10 min to remove unbroken cells and nuclei. The supernatant was loaded on a sucrose cushion [5 ml 35% (wt/vol) sucrose in 10 mM Tris, pH 7.4, with 1 mM EDTA] and centrifuged for 90 min at 150,000 x g at 4 C. The plasma membrane pellets were solubilized for 2 h in 1 ml of digitonin buffer [20 mM Tris-HCl, pH 8.0, 1% (wt/vol) digitonin, 20% (vol/vol) glycerol, 300 mM NaCl, 1 mM EDTA containing 0.1 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin, 5 µ g/ml aprotinin, and 1 µg/ml pepstatin). The 35% sucrose interface (light vesicular fraction) was recovered, diluted in 10 mM Tris, pH 7.4, with 1 mM EDTA and pelleted at 436,000 x g for 90 min in an Optima TL ultracentrifuge (Beckman Coulter, Inc., Fullerton, CA). The light vesicular pellets were subsequently solubilized for 2 h in 1 ml of digitonin buffer.

Receptor Coimmunoprecipitation
Plasma membrane and light vesicular membrane fraction lysates were cleared by centrifugation at 436,000 x g for 20 min at 4 C and incubated (1:500 dilution) with rabbit anti-GFP polyclonal antibody (Molecular Probes, Inc., Eugene, OR) overnight along with 100 µl of a 20% slurry of protein A Sepharose beads (Pharmacia Biotech, Piscataway, NJ) in digitonin buffer plus 3% BSA. The beads were recovered by centrifugation and washed three times in PBS, pH 7.4, and incubated in 200 µl of PBS containing a saturating concentration of 125I-pindolol (~2 nM) at 37 C for 2 h. The beads were recovered by centrifugation and washed three times with PBS pH 7.4. The washed beads were counted in a {gamma}-counter. Nonspecific 125I-pindolol binding to the beads was less than 10% of specific 125I-pindolol binding to ß2AR in the absence of agonist (data not shown).

Agonist-Independent Internalization Assays
The agonist-independent internalization of cell surface receptors was measured by prelabeling cell surface epitope-tagged receptors with primary mouse antiepitope tag antibody (1:500 dilution) on ice for 45 min and then warming cells to 37 C in the absence of agonist for the times indicated in the figure legends. Cells were then transferred back to ice and labeled with the secondary FITC-conjugated antimouse antibody (1:250 dilution) for 45 min. Receptor internalization was defined as the fraction of total cell receptors lost from the cell surface and thus not available to secondary antibodies outside the cell.

Receptor Recycling Assays
Cells expressing epitope-tagged receptors were treated with or without agonist for 30 min at 37 C, washed three times with serum-free media, and either kept on ice or allowed to recover to 37 C for the times indicated in the figure legends. The cells were antibody stained, and the cell surface receptor expression was determined by flow cytometry. Receptor recycling was defined as a recovery of cell surface receptors accessible to antibodies outside the cell after the removal of agonist when compared with the cell surface expression of receptors in matched controls that were not exposed to agonist.

Confocal Microscopy
Confocal microscopy was performed on a LSM-510 laser scanning microscope (Carl Zeiss, Thornwood, NY) using a Zeiss 63x 1.3 NA oil immersion lens. HEK 293 cells expressing ß2AR, AT1AR, and ß2AR/AT1AR chimeras with and without dynaminI-K44A and low levels of either ß-arrestin2-GFP or PKCßII-GFP were plated on 35-mm glass-bottomed culture dishes and kept warm at 37 C in serum-free MEM on a heated microscope stage as described previously (22). Clathrin staining of HEK 293 cells grown on cover slips and fixed with 3.7% paraformaldehyde in PBS with 0.2% Triton X-100 for 20 min was performed using the monoclonal antibody X22 (ATCC) in conjunction with a Texas red-conjugated goat antimouse secondary antibody (Molecular Probes, Inc.). Colocalization studies of ß-arrestin2-GFP and rhodamine-labeled clathrin fluorescence were performed using dual excitation (488, 543 nm) and emission (505–530 nm, GFP; 560 nm, rhodamine) filter sets. Specificity of labeling and absence of signal cross-over were established by examination of single-labeled samples.

Whole-Cell Phosphorylation
Receptor phosphorylation was performed as described previously (38). In brief, the intracellular ATP pool was [32P] labeled by incubating transfected cells seeded in six-well dishes with [32P]orthophosphate (100 µCi/ml) in phosphate- and serum-free medium for 45 min at 37 C. Cells were then treated with or without 1 µM isoproterenol for 10 min at 37 C and washed three times with ice-cold PBS. Resensitized cells were allowed to recover for 20 min at 37 C in agonist-free PBS. The cells were solubilized in radioimmune precipitation buffer (150 mM NaCl, 50 mM Tris, 5 mM EDTA, 10 mM NaF, 10 mM Na2pyrophosphate, 1% NP-40, 0.5% deoxycholate, 0.1% SDS, 0.1 mM phenylmethysulfonyl fluoride, 10 µg/ml leupeptin, 5 µg/ml aprotinin, 1 µg/ml pepstatin A, pH 7.4), and epitope-tagged receptors were immunoprecipitated as described previously (38). In each experiment, equivalent amounts of receptor protein, as determined by receptor expression and the amount of solubilized protein, were subjected to SDS-PAGE followed by autoradiography. The extent of receptor phosphorylation was quantitated using a phosphorimaging system and ImageQuant software (Molecular Dynamics, Inc., Sunnyvale, CA).

Data Analysis
The mean and SEM are expressed for values obtained for the number of separate experiments indicated. Statistical significance was determined by an unpaired two-tailed t test. Time course data were analyzed using PRISM software (GraphPad Software, Inc., San Diego, CA).


    ACKNOWLEDGMENTS
 
We would like to thank Drs. S. A. Laporte, M. G. Caron, and D. J. Kelvin for helpful discussions and critical reading of the manuscript.


    FOOTNOTES
 
Address requests for reprints to: Stephen S. G. Ferguson, Robarts Research Institute, 100 Perth Drive, P.O. Box 5015, London, Ontario, Canada N6A 5K8. E-mail: ferguson{at}rri.on.ca

P.H.A. is the recipient of a Merck Frosst Canada Postdoctoral Fellowship. L.D. is the recipient of a John P. Robarts Research Institute studentship. S.S.G.F. is the recipient of a McDonald Scholarship Award from the Heart Stroke Foundation of Canada. This work was supported by Heart and Stroke Foundation of Ontario Grants NA-3349 and T 4047 and Medical Research Council of Canada Grant MA-15506.

1 Abbreviations used in this paper: AT1AR, angiotensin type 1A receptor; AT1AR-ß2CT, AT1AR-based chimera with the ß2AR carboxyl-terminal tail; ß2AR, ß2-adrenergic receptor; ß2AR-ATCT, ß2AR-based chimera with the AT1AR carboxyl-terminal tail. Back

Received for publication May 5, 2000. Revision received August 9, 2000. Accepted for publication August 23, 2000.


    REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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