Parathyroid Hormone Receptor Recycling: Role of Receptor Dephosphorylation and ß-Arrestin
Stephanie Chauvin,
Margaret Bencsik,
Tom Bambino and
Robert A. Nissenson
Endocrine Research Unit, Veterans Affairs Medical Center, University of California San Francisco, San Francisco, California 94121
Address all correspondence and requests for reprints to: Dr. R. A. Nissenson, Endocrine Research Unit (111N), 4150 Clement Street, San Francisco, California 94121. E-mail: chicago{at}itsa.ucsf.edu.
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ABSTRACT
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The recovery of PTH receptor (PTHR) function after acute homologous receptor desensitization and down-regulation in bone and kidney cells has been attributed to receptor recycling. To determine the role of receptor dephosphorylation in PTHR recycling, we performed morphological and functional assays on human embryonic kidney 293 cells stably expressing wild-type (wt) or mutant PTHRs. Confocal microscopy and ligand binding assays revealed that the wt PTHR is rapidly recycled back to the plasma membrane after removal of the agonist. Receptors that were engineered to either lack the sites of phosphorylation or to resemble constitutively phosphorylated receptors were able to recycle back to the plasma membrane with the same kinetics as the wt PTHR. The PTHR was found to be dephosphorylated by an enzyme apparently distinct from protein phosphatases 1 or 2A. The PTHR and ß-arrestin-2-green fluorescent protein (GFP) were found to stably colocalize during PTHR internalization, whereas after agonist removal and during receptor recycling, the colocalization slowly disappeared. Experiments using phosphorylation-deficient PTHRs and a dominant-negative form of ß-arrestin showed that ß-arrestin does not regulate the efficiency of PTHR recycling. These studies indicate that, unlike many G protein-coupled receptors, PTHR recycling does not require receptor dephosphorylation or its dissociation from ß-arrestin.
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INTRODUCTION
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THE PTH RECEPTOR (PTHR) is a class II G protein-coupled receptor (GPCR) that transduces the biological responses to PTH and to PTHrP. The PTHR is the major receptor responsible for initiating the pleiotropic paracrine effects of PTHrP as well as the endocrine actions of PTH on calcium and skeletal homeostasis (1). To control the magnitude and the timing of signaling by GPCRs, receptors are subject to a variety of regulatory processes including desensitization, internalization, and down-regulation. Receptor desensitization limits the magnitude and duration of receptor signaling by uncoupling the receptor from its cognate G protein, whereas internalization results in translocation of the receptor from the plasma membrane to the endocytic pathway of the cell. Once a GPCR has been desensitized and internalized, it may be either directed to lysosomes for degradation (down-regulation), or recycled back to the plasma membrane in an active form (resensitization).
Previous studies have demonstrated that the PTHR is rapidly phosphorylated by a GPCR kinase (GRK) upon agonist stimulation (2, 3, 4, 5) and then internalized via clathrin-coated pits (6). These events are associated with rapid desensitization and down-regulation of the PTHR in target cells (3). However, the post-endocytic trafficking pathways of the PTHR are still unclear. To better understand how PTH responsiveness is regulated, it is crucial to determine whether the PTHR recycles back to the plasma membrane or undergoes lysosomal degradation after endocytosis. Previous investigations on bone and kidney cells showed a recovery of receptor function after initial desensitization due to acute PTH stimulation and suggested a role for PTHR recycling in resensitization (7, 8). Recycling of the PTHR is therefore likely to be an important mechanism for the maintenance of appropriate responsiveness to PTHR agonists (PTH and PTHrP) under physiological circumstances. However, direct studies of the recycling of the PTHR to the plasma membrane are lacking.
Much of our knowledge concerning the molecular basis of GPCR recycling is derived from studies on the ß-adrenergic receptor, a class I GPCR. Exposure of cells to ß-adrenergic agonists leads to rapid receptor phosphorylation, desensitization, and internalization via clathrin-coated pits (9). Within about 1 h, the receptor appears in early recycling endosomes that contain phosphatases required for receptor GRK dephosphorylation, a critical step in the ß-adrenergic receptor resensitization pathway (10, 11, 12, 13). The V2 vasopressin receptor is also a model of class I GPCR, which recycles rapidly to the plasma membrane, but only once it has been dephosphorylated (14). How dephosphorylation promotes receptor recycling is unclear. Nonetheless, it is evident for these class I GPCRs that phosphorylation regulates not only receptor-G protein coupling but also the subsequent trafficking of the GPCR in the endocytic pathway.
Recently, evidence has accumulated that ß-arrestin also participates in the recycling of some GPCRs. Activation of almost all GPCRs induces translocation of ß-arrestin from the cytosol to the plasma membrane (15, 16). A subset of GPCRs (e.g. the vasopressin V2 receptor) remain associated with ß-arrestin during the process of endocytic trafficking of the receptor. The stability of this association has been suggested to inhibit receptor recycling, thereby preventing receptor resensitization (17). In the case of the PTHR, receptor activation induces ß-arrestin translocation to the plasma membrane (6, 18). Binding of arrestin to the PTHR apparently inhibits PTHR-G protein coupling, thereby leading to termination of receptor signaling (19).
In the present study, we investigated the post-endocytic trafficking of a prototype class II GPCR (the PTHR), and particularly whether phosphorylation of the receptor and/or its association with ß-arrestin regulates receptor trafficking or recycling. The results demonstrate directly that the PTHR recycles to the plasma membrane after endocytosis, and little if any of the receptor as targeted for degradation. Recycling of the PTHR is not modulated by receptor phosphorylation. Furthermore, the PTHR is stably associated with ß-arrestin after initial endocytosis, but this strong association does not inhibit the ability of the PTHR to recycle to the plasma membrane. Thus, in the case of this class II GPCR, the major role of receptor phosphorylation and arrestin binding appears to be to regulate the early steps in receptor desensitization and internalization.
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RESULTS
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Down-Regulation of the PTHR Is Not Associated with PTHR Degradation
Although a number of studies have demonstrated functional down-regulation of the PTHR after exposure of target cells to agonists, the extent to which this results from post-endocytic degradation of the PTHR is unclear. In previous studies, we have shown that PTHR internalization is extensive and rapid in human embryonic kidney (HEK)-wild type (wt) cells after the addition of PTH (20). In the present study, we found a 60% reduction in PTHR activity after a 30-min exposure of these cells to PTH (Fig. 1A
). Longer exposure of cells to PTH (up to 24 h) had little additional effect on PTHR activity. These experiments were carried out under conditions where the level of specific ligand binding was proportional to the number of receptors (21), suggesting that the decrease reflects a change in the number of PTHRs. Previous studies in bone and kidney cells have demonstrated a marked down-regulation of functional PTHRs, an effect suggested to result from lysosomal degradation of the receptor. However, Western blots of whole cell lysates from HEK-wt cells demonstrated that the level of intact PTHR was unchanged by exposure to PTH (Fig. 1
, B and C). Maintenance of PTHR levels was obtained in the presence of cycloheximide, indicating that de novo synthesis of the receptor was not required to maintain cellular levels of the PTHR. No evidence for degradation of the PTHR was obtained even after prolonged treatment of HEK-wt cells with PTH, strongly indicating that the PTHR is functionally down-regulated without a significant reduction in total receptor protein.

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Figure 1. Effect of PTH Treatment on Total Cellular Expression of the PTHR
HEK-wt PTHR cells were treated at 37 C for 30 min with or without 1 µM bPTH(134). At the end of the incubation, the cells were subjected to radioligand binding, as described in Materials and Methods (A). HEK-wt PTHR cells were treated at 37 C for either 3 or 24 h in the absence (B) or 3 h in presence of cycloheximide (C) with or without 1 µM bPTH(134). Cells were solubilized and cellular proteins were resolved by SDS-PAGE, and Western blots of the PTHR were carried out as described in Materials and Methods. Each lane was loaded with equivalent amounts of total cell protein. The exposed films were scanned and quantified using Bio-Rad Laboratories, Inc. Molecular Analyst Program. Data represent the mean ± SE of three or more independent experiments.
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Internalized PTHRs Rapidly Recycle to the Plasma Membrane after Removal of Agonist
GPCRs vary considerably in their recycling behavior. Some like the thrombin receptor display little or no recycling to the plasma membrane, but rather are directly trafficked to lysosomes for degradation (22). Others, like the type 1A angiotensin II receptor, recycle relatively slowly (within hours of removal of agonist) (23). In contrast, the ß-adrenergic receptor is efficiently recycled to the plasma membrane within about 1 h after the removal of agonist (24). Therefore, it was of interest to determine whether the PTHR recycles back to the plasma membrane after internalization and to assess the kinetics of this process.
To visualize the trafficking of the PTHR to and away from the cell surface, we used a morphological approach employing laser confocal microscopy. Immunostaining of the PTHR demonstrated that the receptor is predominantly localized to the plasma membrane in the basal state, and that it rapidly translocates into intracellular vesicles after the addition of 1 µM bovine (b) PTH(134) (Fig. 2
). After acid washing to remove PTH from the cell surface, the cells were either fixed immediately or incubated at 37 C (up to 4 h) before fixation. The reappearance of receptors at the cell surface after agonist treatment was rapid, requiring only about 1 h to recover to a new steady state in which the cell-surface receptor density was close to that seen in untreated cells. These experiments were done in the presence of cycloheximide indicating that this recovery resulted from receptor recycling rather than from the biosynthesis of new receptor protein. We also observed that PTHR recycling required removal of the agonist and was not evident in the continuous presence of bPTH(134), even after several hours (data not shown).

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Figure 2. PTH-Induced Endocytosis and Rapid Recycling of the PTHR
HEK-wt PTHR cells were fixed and immunostained using a mouse antiopossum PTHR monoclonal antibody and fluorescein isothiocyanate-conjugated donkey antimouse IgG. Micrographs (magnification, x100) show PTHR immunolocalization on the cell membrane in untreated cells (basal), internalization after 30 min incubation with 1 µM bPTH(134) (PTH), and relocalization to the cell membrane rapidly after removal of the ligand (30 min, 1 h, 2 h, and 4 h recycling). All experiments were performed in the presence of cycloheximide. The results are representative of those obtained in five separate experiments.
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As a second approach for assessing PTHR recycling, we used a radioligand binding assay to evaluate the presence of functional, cell surface PTHRs. In this approach, HEK-wt PTHR cells were treated with a saturating concentration of PTH to induce receptor endocytosis. PTH was then removed from the medium, and the recovery of cell surface PTHR was evaluated by binding assay at various times of recycling. As shown in Fig. 3
, the time course for recycling of the PTHR was slightly different from that seen with confocal microscopy. The functional assay demonstrated that about 50% of the cell surface PTHRs persisted after PTH treatment, and that recycling of functional receptors could be detected as early as 30 min after the removal of PTH. Full recycling of functional PTHRs was evident within about 1 h. Similar results were obtained in the presence and absence of cycloheximide (data not shown). The basis for the small differences in recycling kinetics as revealed by functional assay vs. confocal microscopy is probably due to the greater sensitivity and quantitative nature of the functional assay.

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Figure 3. Functional Recycling of the PTHR
HEK-wt PTHR cells were incubated with or without 1 µM bPTH(134) for 30 min at 37 C (+PTH or basal, respectively), followed by removal of the agonist and further incubated at 37 C to allow receptor recycling for the times indicated. These experiments were performed in the presence of cycloheximide to exclude the possible contribution of new receptor synthesis. The number of receptors present at the cell surface was measured by radioligand binding and expressed as the percentage of specific binding. Data represent the mean ± SE of nine independent experiments.
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Relationship between the Phosphorylation State of the PTHR and Recycling
Agonist binding to the ß-adrenergic receptor results in receptor phosphorylation and internalization. Subsequently, the receptor is dephosphorylated in an acidified vesicular compartment by protein phosphatase 2A. After dephosphorylation, the receptor recycles back to the plasma membrane, and resensitization is achieved (10, 11, 12, 13). We have previously reported that the PTHR is GRK phosphorylated after agonist binding (5), but the regulation of dephosphorylation of the PTHR and its role in receptor recycling is unclear. Figure 4
illustrates the time course of PTHR dephosphorylation and shows that the removal of agonist resulted in receptor dephosphorylation with kinetics very similar to those observed for PTHR recycling. After approximately 1 h, the amount of 32P incorporated into the receptor had returned to basal levels.

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Figure 4. Kinetics of PTHR Dephosphorylation
HEK-wt PTHR cells were labeled for 2 h in phosphate-free media containing 100 µCi [32P]-orthophosphoric acid at 37 C. Cells were treated for 30 min at room temperature with 240 nM bPTH(134). After removal of the agonist by acid washes, cells were incubated at 37 C for the time indicated and then solubilized. PTHRs were immunoprecipitated and resolved by SDS-PAGE. 32P incorporation was quantified from autoradiograms using the Bio-Rad Laboratories, Inc. GS-363 Phosphorimager system. Results are expressed as the increase in phosphorylation of the PTHR in cells exposed to PTH (minus basal phosphorylation), expressed as a percentage of that seen at the time of removal of PTH. Results are the mean ± SE of six independent experiments.
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Studies on the ß-adrenergic receptor have shown that okadaic acid or calyculin A, both potent inhibitors of protein phosphatase 1 (PP1) and 2A (PP2A) (25), inhibited ß-adrenergic receptor dephosphorylation and thereby its recycling (10, 13, 26). Because the kinetics of recycling of the ß-adrenergic receptor and the PTHR are very similar, we investigated whether okadaic acid or calyculin A were also able to prevent functional recycling of the PTHR. Figure 5
shows that PTHR recycling was not affected by the presence of high doses of either okadaic acid or calyculin A. Similarly, these agents had no detectable effect on the rate of recycling of the PTHR after removal of PTH when assessed by confocal microscopy (data not shown). Striking, neither okadaic acid nor calyculin A, at doses that are efficacious in blocking dephosphorylation of the ß-adrenergic receptor, had any affect on the dephosphorylation of the PTHR after removal of PTH (Fig. 6
). These findings suggest that the regulation of dephosphorylation of the PTHR is mediated by a phosphatase other than PP1 or PP2A, and may also indicate that the ability of the PTHR to efficiently recycle to the plasma membrane is not dependent upon receptor dephosphorylation.

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Figure 5. Effect of Okadaic Acid and Calyculin A on PTHR Recycling
The effects of 1 µM okadaic acid or 2 nM calyculin A on PTHR recycling was assessed in HEK-wt PTHR cells, using the procedure described in Fig. 3 . Results are the mean ± SE of five independent experiments with okadaic acid, and four independent experiments with calyculin A.
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Figure 6. Effect of Okadaic Acid on PTHR Dephosphorylation
Dephosphorylation experiments were performed on HEK-wt PTHR cells using the procedure described in Fig. 4 , in the absence or continuous presence of 1 µM okadaic acid. Results are the mean ± SE of three independent experiments.
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To further explore the relationship between PTHR phosphorylation and recycling, we investigated the recycling of a PTHR mutated in the cytoplasmic tail to eliminate the sites of agonist-stimulated phosphorylation. The mutated PTHR (S(483498)A) resembles the wt PTHR in its ability to bind PTH and activate adenylyl cyclase but displays no detectable phosphorylation in response to agonist treatment (5). We also studied the trafficking of a mutated PTHR in which the serine residues that are the targets of agonist-dependent phosphorylation were replace with acidic residues. This mutant PTHR (S(483498)D/E) is expected to display properties of a constitutively-phosphorylated PTHR, and represents a strategy successfully used by others to investigate the effects of protein phosphorylation (27, 28, 29). Functional analysis of the S(483498)D/E PTHR demonstrated that it retains the ability to bind bPTH(134) with the same affinity as the wt (dissociation constant = 5.34 nM vs. dissociation constant = 9.66 nM) and to signal normally (EC50= 35.5 pM vs. EC50= 22.4 pM) (Table 1
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Morphological (Fig. 7A
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) studies performed in the presence of cycloheximide demonstrated that the recycling of the S(483498)A mutant was similar to that of the wt PTHR, with recycling to the plasma membrane evident within 1 h after removal of PTH. The S(483498)D/E mutant was also efficiently recycled to the cell surface. Taken together, these results indicate that in contradistinction to other GPCRs like the V2 vasopressin and ß-adrenergic receptors, phosphorylation is not a major factor in the regulation of recycling of the PTHR.

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Figure 7. Role of PTHR Phosphorylation in Recycling
Recycling was studied for PTHRs that either lack phosphorylation sites (S(483498)A) or contain acidic residues to mimic phosphorylation (S(483498)D/E). These mutant PTHRs were stably expressed in HEK293, and recycling was examined after PTH-induced internalization, using the procedure described in Fig. 2 (A) and Fig. 3 (B) . In A, the PTHRs were visualized using a donkey antimouse IgG-fluorescein-conjugated secondary antibody (S(483498)D/E) or a goat antimouse IgG-rhodamine conjugated secondary antibody (S(483498)A) followed by confocal microscopy. In B, the percent recycling was calculated as follows: [(bound ligand after recycling - bound ligand before recycling)/(internalized ligand)] x 100. All experiments were performed in the presence of cycloheximide. Data represent the mean ± SE of three independent experiments.
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Role of ß-Arrestin in the Recycling of the PTHR
Virtually all GPCRs interact with arrestins after agonist binding, but recently it has become clear that the fate of the GPCR-ß-arrestin complex is not uniform (30). For some GPCRs (e.g. ß-adrenergic receptors), the receptor-ß-arrestin complex dissociates at or near the plasma membrane shortly after formation of clathrin-coated pits. For other GPCRs (e.g. vasopressin V2 receptors), the receptor-ß-arrestin complex is more stable, and the proteins are internalized into endosomes as a complex.
Whereas the ß-adrenergic receptor dissociates from ß-arrestin and recycles to the membrane within 1 h (23, 24), we recently showed that the PTHR cointernalizes with ß-arrestin during PTH treatment, a process that is facilitated by receptor phosphorylation (18). In the present study, we assessed whether PTHR recycling requires dissociation of the receptor-ß-arrestin-2 complex. As shown in Fig. 8
, addition of PTH to cells expressing the wt PTHR resulted in rapid translocation of ß-arrestin-2-green fluorescent protein (GFP) from the cytosol to the plasma membrane and subsequent colocalization with the PTHR in endocytic vesicles. Parallel experiments with the S(483498)A PTHR demonstrated a more transient interaction between the receptor and ß-arrestin-2-GFP even in the continued presence of PTH. The t1/2 for dissociation of ß-arrestin-2 from the S(483498)A PTHR was about 15 min (not shown). This is faster than the t1/2 for recycling of the S(483498)A PTHR receptor, making it unlikely that ß-arrestin binding is required for efficient PTHR recycling. After removal of the agonist and during receptor recycling, the colocalization between the wt PTHR and ß-arrestin-2-GFP slowly disappeared. Some receptors remained colocalized with ß-arrestin-2-GFP even 1 h after removal of PTH.

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Figure 8. Colocalization of ß-Arrestin2-GFP with Internalized wt and S(483498)A Mutant PTHRs
HEK-wt PTHR and HEK-S(483498)A PTHR cells were transiently transfected with a plasmid containing the cDNA for ß-arrestin-2-GFP. Cells were treated or not (basal) with 1 µM bPTH(134) for the indicated times, fixed, and immunostained with a mouse antiopossum PTHR monoclonal antibody and rhodamine-conjugated goat antimouse IgG. Another set of cells was treated for 30 min with 1 µM bPTH(134), acid washed, and incubated at 37 C for 30 min, 1 h, or 4 h to allow receptor recycling. Shown are representative confocal microscopy images of PTHR receptor immunofluorescence (red) and ß-arrestin-2-GFP fluorescence (green). Colocalization (yellow) of the receptor with ß-arrestin-2-GFP is shown in the overlay. All experiments were performed in the presence of cycloheximide.
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To determine whether ß-arrestin is a negative regulator of PTHR recycling, we overexpressed a dominant-negative form of ß-arrestin (ß-arrestin-1-V54D) in HEK-wt PTHR cells. This dominant negative arrestin has been shown to inhibit the internalization of other GPCRs such as the ß-adrenergic receptor, the endothelin receptor, and the
-opioid receptor (31, 32, 33). If endogenous ß-arrestin is a negative regulator of receptor recycling, we would expect to observe a faster receptor recycling in cells expressing dominant-negative arrestin. As shown in Fig. 9
, expression of ß-arrestin-1-V54D failed to alter the rate of recycling of the wt PTHR. This result was obtained despite the fact that ß-arrestin-1-V54D was well expressed (by Western blotting) and inhibited the agonist-induced endocytosis of the PTHR. Similarly, transient cotransfection of the PTHR and ß-arrestin-1-V54D in 293 cells resulted in agonist-induced PTHR recycling that was indistinguishable from recycling observed after transient expression of the PTHR alone (n = 5, data not shown). Taken together, these results suggest that arrestin does not directly regulate the recycling of the PTHR.

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Figure 9. Role of ß-Arrestin in PTHR Recycling
HEK293 cells transiently transfected with the wt PTHR alone or with a dominant-negative (dn) form of ß-arrestin-1 (ß-arrestin-1-V54D) were treated with bPTH(134) for 30 min (+PTH). After washing out the agonist, the cells were incubated at 37 C to allow recycling for the times indicated. The cells were then fixed, immunostained, and observed by confocal microscopy as described in Materials and Methods (A). The results are representative of those obtained in three separate experiments. In some cases, cells were solubilized and resolved by SDS-PAGE, transferred and blotted for ß-arrestin (B). HEK293 cells transiently transfected with the wt PTHR alone or with ß-arrestin-1-V54D (dn-arr1) were treated with 1 µM bPTH(134), and the internalization was measured after 10 min of exposure to [125I]PTHrP as described in Materials and Methods (B). Data represent the mean ± SE of three or more independent experiments. *, P < 0.01. All experiments were performed in the presence of cycloheximide.
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DISCUSSION
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Once internalized, most GPCRs are recycled back to the plasma membrane, but some are routed to lysosomes and targeted for degradation. In the case of the ß-adrenergic receptor, known to be sorted to the recycling pathway, internalization to the endosomal compartment results in ligand dissociation followed by receptor dephosphorylation. Receptor dephosphorylation is performed by a membrane-associated phosphatase that resides in an acidic endosomal compartment and has specificity for GRK-phosphorylated GPCRs (13, 26). After dephosphorylation, the receptor is recycled to the plasma membrane as a fully functional receptor (9, 34). It has been suggested that the rate of receptor dephosphorylation is dependent on the rate of dissociation of arrestin from the phosphorylated receptor (17). Consistent with this, the ß-adrenergic receptor rapidly dissociates from ß-arrestin early during the internalization process, and this is associated with rapid dephosphorylation and resensitization of the receptor after the removal of agonist (17). Conversely, the V2 vasopressin receptor is internalized as a stable complex with ß-arrestin, and this is associated with much slower resensitization of the receptor (17).
One goal of the present study was to determine the fate of the PTHR after agonist-induced endocytosis. We observed that the wt PTHR expressed in 293 cells is susceptible to functional down-regulation but that this is not associated with degradation of the receptor even after prolonged exposure to PTH. These findings indicate that the PTHR is maintained in an intact, potentially functional form within the endocytic compartment during continuous exposure to agonist. Removal of PTH results in nearly complete recycling of the PTHR to the plasma membrane within approximately 1 h. Ferrari et al. (6) have reported that the human PTHR recycles to the plasma membrane incompletely even 2 h after removal of PTH under similar conditions to those used in our study (i.e. in the presence of cycloheximide to block de novo receptor synthesis). The basis for the difference is unclear, but may relate to the different species of PTHR studied (opossum vs. human). However, our data are in accordance with previous results using indirect methods in bone and kidney cells suggesting recycling of the PTHR within 12 h after removal of the ligand (7, 8). Our findings indicate that down-regulation of the cell surface PTHR after chronic exposure to agonists can be attributed to relocation of the intact receptor to an intracellular pool, and that the receptor can be rapidly recalled to the plasma membrane in a functional form as the concentration of extracellular agonist falls.
An additional objective of this study was to assess whether the phosphorylation state of the PTHR is an important determinant of PTHR recycling. Previous studies have shown that phosphorylation of the PTHR occurs exclusively on serine residues in the proximal portion of the cytoplasmic tail (2, 3, 5). We evaluated the recycling of two phosphorylation mutants of the PTHRone lacking all of the phosphorylation sites (S(483498)A), and one in which these serine residues were replaced by acidic amino acids (S(483498)D/E) to mimic the effect of agonist-dependent phosphorylation. Functional and morphological studies revealed that these two mutant receptors were able to recycle back to the cell surface after agonist-mediated endocytosis, with the same kinetics as observed for the wt PTHR. Thus, the efficiency of recycling of the PTHR (unlike the ß-adrenergic receptor) is not detectably influenced by the phosphorylation state of the receptor. Nonetheless, the PTHR is clearly dephosphorylated after the removal of agonist, and this process occurs with a time course resembling that of receptor recycling. PTHR dephosphorylation was insensitive to okadaic acid and calyculin A, suggesting that PP1 and PP2A are not the relevant phosphatases. Similar results have recently been reported for dephosphorylation of the D1 dopamine and V1a vasopressin receptors (35, 36). Because these findings with okadaic acid and calyculin A represent negative data, more extensive studies will be needed to identify the nature of the phosphatase responsible for dephosphorylation of the PTHR. Interestingly, the D1 dopamine receptor is rapidly dephosphorylated after internalization but does not recycle efficiently, suggesting that the model depicted for the ß-adrenergic receptor correlating receptor dephosphorylation to recycling may not be applicable to all GPCR or to all cell types. The results indicate that certain GPCRs are susceptible to dephosphorylation by a novel phosphatase, whose action does not target the receptor to the plasma membrane but rather allows for effective signaling once the receptor is recycled.
Finally, we examined the role of ß-arrestin in PTHR recycling. Previous studies have demonstrated that the activated PTHR recruits ß-arrestin from the cytosol to the plasma membrane, and that the PTHR is internalized with ß-arrestin (18). In the present study, we visualized colocalization between ß-arrestin-2-GFP and the PTHR during receptor endocytosis, whereas the proteins became dissociated during the course of PTHR recycling. Dissociation of the PTHR and ß-arrestin-2-GFP occurred with a time course similar to that of PTHR dephosphorylation, a finding consistent with the reported role of PTHR phosphorylation in promoting the stable interaction between the PTHR and ß-arrestin (18). It is likely that dissociation of ß-arrestin is a prerequisite for dephosphorylation of the PTHR by the relevant phosphatase, although further studies are required to establish this point. Two lines of evidence suggest that association of the PTHR with ß-arrestin does not have a major impact on PTHR recycling. First, the phosphorylation-deficient PTHR mutant that dissociates readily from ß-arrestin after endocytosis did not display recycling kinetics that differed detectably from those displayed by the wt PTHR. Secondly, expression of a dominant-negative form of ß-arrestin did not alter the efficiency of PTHR recycling. It is unclear whether the stable association of ß-arrestin with the internalized PTHR is functionally important. It is conceivable that this association is required to maintain the desensitized state of the PTHR in intracellular compartments or to initiate and/or regulate other signaling pathways. With respect to the latter possibility, recent studies have demonstrated that ß-arrestin-bound, internalized GPCRs may initiate activation of MAPK (37). A similar mechanism could account for the reported ability of the PTHR to mediate activation of MAPK (38).
Using chimeric receptors, it has been shown that the C-terminal tail of GPCRs can direct the intracellular fate of the receptor (22). The C-terminal tails of some GPCRs have a great influence on the sorting of these internalized GPCRs to either the degradation or recycling pathways (17, 39). Cao and colleagues (40) showed that the interaction of the C-terminal DSLL sequence of the ß-adrenergic receptor with the PDZ domains of EBP50 is necessary for the recycling of the internalized ß-adrenergic receptor. Moreover, Kishi and co-workers (41) have proposed that ligand binding may be an important determinant of the fate of internalized receptors. Accordingly, it is plausible that there is a specific motif in the C-terminal tail of the PTHR that directs the receptor to the recycling pathway rather than to a degradative fate. Further studies are in progress to determine the precise location of the sorting motifs that route the internalized PTHR to recycling pathway.
In summary, the present study demonstrates that, after activation and internalization, the PTHR is rapidly recycled back to the cell surface rather than degraded. This recycling process is independent of receptor dephosphorylation. The results further indicate that while the PTHR is internalized stably complexed with ß-arrestin, maintenance of this complex does not impair the recycling of the PTHR. The ability of the PTHR to be maintained in an intact state after internalization, and to be rapidly recruited in a functional form to the plasma membrane are undoubtedly critical to the recovery of responses to PTH and PTHrP after exposure of target cells to high levels of these agonists in vivo.
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MATERIALS AND METHODS
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Materials
bPTH(134) and PTHrP(134) were obtained from Bachem (Torrance, CA). HEK293 cells and the expression vector pCEP4 were obtained from Invitrogen (San Diego, CA). Restriction enzymes were purchased from New England Biolabs, Inc. (Beverly, MA). Oligonucleotides were synthesized and purified at the University of California San Francisco (UCSF) Biomolecular Resource Facility. Rat V53D-ß-arrestin-1 and ß-arrestin-2-GFP in pcDNA.1 were gift from Dr. M.G. Caron (Duke University, Durham, NC). Polyclonal ß-arrestin antibody was provided by Dr. Michael Bliziotes [Veterans Affairs Medical Center (VAMC), Portland, OR]. Confocal microscopy was performed at the VAMC-SF Imaging Core Facility.
Generation of PTHR and PTHR Mutant Expressing Cell Lines
Site-directed mutagenesis was performed using the Transformer Site-Directed Mutagenesis Kit (CLONTECH Laboratories, Inc., Palo Alto, CA) on the opossum PTH/PTHrP receptor cDNA in pBluescript II SK (Stratagene, La Jolla, CA). The S(483498)A PTHR mutant was generated and subcloned into pcDNA 3.1 as previously described (5). The mutagenic primer for the S(483498)D/E mutant was designed to replace Serine residues 483, 485, 486, 489, 495, and 498 (483SGSSTYSYGPMVSHTS498) with an acidic residue (483DGDDTYDYDPMVEHTD498). An SmaI restriction site was also introduced to aid in the identification of the mutant clone. The mutagenic primer sequence is as follows:
SmaI 483 485 486 489
5' GG AAG GCC CGG GAT GGC GAC GAT ACC TAC GAC TAT GGC CCC ATG
495 498
GTG GAA CAT ACA GAT GTC ACC AAT GTG 3'
Oligonucleotides were synthesized and purified by the UCSF Biomolecular Resource Center. After generation of the S(483498)D/E mutant receptor, the cDNA was subcloned into a mammalian expression vector pCEP4 using restriction sites HindIII and NotI.
Cell Culture and Transfection
HEK293 cells were maintained in DMEM containing 10% fetal calf serum and antibiotics (100 U/ml penicillin and 100 µg/ml streptomycin) in 7.5% CO2 at 37 C. Calcium phosphate-mediated transfection was performed as previously described (42). After overnight incubation at 37 C, cells were washed twice with calcium- and magnesium-free PBS, grown for another day in DMEM, and then subjected to selection with 200 µg/ml Hygromycin B for at least 3 wk.
Localization of the PTHR in Fixed Cells by Direct Immunofluorescence
HEK293 cells stably expressing the wt (HEK-wt) or mutant (HEK-mutant: HEK-S(483498)A and HEK-S(483498)D/E) PTHRs were grown on glass coverslips. The cells were pretreated with 25 µg/ml cycloheximide in DHB (20 mM HEPES, 0.1% BSA in DMEM) for 1 h at 37 C, and then treated with 1 µM bPTH(134) for 30 min at 37 C, transferred to ice, rinsed with PBS, and then fixed with 4% paraformaldehyde for 20 min. For recycling time points, PTH was removed by washing the cells with 50 mM glycine, 100 mM NaCl (pH 3.0), and then incubated in DHB containing cycloheximide (DHBC) at 37 C for the time indicated in the figure legends. Once the cells were fixed and washed three times with PBS, they were permeabilized and blocked using a 0.2% Triton X-100, 1% dry milk, 0.3% cold fish gelatin solution for 1 h at room temperature. Affinity-purified monoclonal primary antibody was diluted 1:500 and applied to the specimens incubating overnight at 4 C. This primary antibody is directed against the receptors first extracellular domain (43). After three PBS washes, donkey fluorescein isothiocyanate or goat Rhodamine-tagged secondary antibody (antimouse IgG, Santa Cruz Biotechnology, Inc., Santa Cruz, CA) was diluted 1:200 and applied to the cells for 90 min at room temperature. Coverslips were then rinsed with PBS and mounted for confocal microscopy (Leica Corp., Exton, PA; model TCS NT/SP).
Assay for Recovery of Receptors to the Cell Surface
HEK-wt or HEK-mutant PTHRs cells diluted with HEK293 cells (1/3) were grown for 5 d in 35-mm wells to confluence. The cells were preincubated at 37 C for 1 h in DHBC and then treated with 1 µM bPTH(134) for 30 min at 37 C. After two washes with ice-cold PBS, bPTH(134) was washed out with 2-fold 5-min incubations on ice with 50 mM glycine, 100 mM NaCl (pH 3.0). The cells were washed three times and then incubated in DHBC for various times at 37 C. At the end of each incubation period, a binding assay was performed at 4 C for 2 h with 200,000 cpm 125I-PTHrP in the presence or absence of 1 µM bPTH(134) as described below.
Assays for Receptor Function
Ligand binding and cAMP assays were carried out after our previously published procedures (5). For binding studies, HEK-wt or HEK-mutant PTHRs were incubated in DHB for 2 h on ice, followed by a 2-h incubation in the same buffer containing 200,000 cpm 125I-PTHrP(134) with or without unlabeled bPTH(134). The cells were washed twice with PBS and extracted with 0.8 N NaOH. Cell-associated 125I-PTHrP was then counted. Competitive curves were fitted to a one site competitive binding curve. To measure the cAMP levels, the cells were incubated in DHB containing 1 mM isobutylmethylxanthine and a various concentrations of bPTH(134) for 10 min at room temperature. After two washes with ice-cold PBS, the cellular cAMP was extracted with 95% ethanol and quantified by RIA.
Down-Regulation Assay
The down-regulation assay was performed by assessing the observed change in the total number of receptors determined through Western blotting, and by the binding of 125I-PTHrP, as described above. HEK-wt PTHR cells were grown for 3 d in 35-mm wells and preincubated in DHB for 1 h at 37 C. The cells were incubated in DHB with or without 1 µM bPTH(134) for 30 min. The cells were then washed with ice-cold PBS and subjected to radioligand binding with saturating concentration of 125I-PTHrP. With another set of cells treated in the same way as those just described, equal amount of cell lysate solubilized in RIPA buffer (1% Nonidet P-40; 0.5% deoxycholate; 0.1% sodium dodecyl sulfate; 50 mM Tris, pH 7.4; 100 mM NaCl; 2 mM EDTA; 50 mM NaF) were resolved by SDS-PAGE (7.5% acrylamide), transferred to nitrocellulose membranes, and blotted for PTHR using OK1 monoclonal antibody followed by detection using horseradish peroxidase-conjugated sheep antimouse antibody and electrochemiluminescence (Amersham Pharmacia Biotech, Arlington Heights, IL). Immunoblots were quantified by densitometric scanning of films exposed in the linear range.
Measurement of Receptor Dephosphorylation
To measure phosphorylation/dephosphorylation of the PTHR in intact cells, HEK-wt PTHR cells were labeled in six-well plates with 100 µCi/well [32P]orthophosphate in 25 µg/ml cycloheximide, phosphate-free DMEM (DMEMPC) for 2 h. The cells were treated with 1 µM bPTH(134) for 30 min at 37 C in DMEMPC. After two washes with ice-cold PBS and two 5-min incubations on ice with 50 mM glycine, 100 mM NaCl (pH 3.0), the cells were washed three times and incubated in DMEMPC for various times at 37 C. The cells were then solubilized in RIPA buffer supplemented with phosphatase inhibitors (300 nM okadaic acid, 10 mM tetrasodium pyrophosphate, 0.1 mM sodium orthovanadate, and 10 mM sodium fluoride) for 1 h on ice, and the receptors were immunoprecipitated with 10 µg of mouse monoclonal antiopossum PTHR antibody immobilized on Sepharose beads. Immunoprecipitated proteins were analyzed by SDS-PAGE and Phosphorimager (Bio-Rad Laboratories, Inc., Hercules, CA) analysis.
Colocalization of wt and S(483498)A PTHRs and ß-Arrestin 2-GFP
HEK-wt and S(483498)A PTHR cells were grown on glass coverslips and transfected with a plasmid encoding ß-arrestin 2 fused to EGFP (ß-arrestin2-GFP). Forty-eight hours after transfection, the cells were incubated with 1 µM bPTH(134) for 30 min at 37 C, acid-washed, and incubated at 37 C for different times of recycling. The cells were then fixed, immunostained, and observed by confocal microscopy, as described above.
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FOOTNOTES
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This work was supported by "la Ligue contre le Cancer" (to S.C.), by NIH Grant DK-35323 (to R.A.N.) and by a Merit Review grant from the Department of Veterans Affairs (to R.A.N.). R.A.N. is a Senior Research Career Scientist of the Department of Veterans Affairs.
Abbreviations: b, Bovine; DMEMPC, phosphate-free DMEM; GFP, green fluorescent protein; GPCR, G protein-coupled receptor; GRK, GPCR kinase; HEK, human embryonic kidney; PP, protein phosphatase; PTHR, PTH receptor; wt, wild-type.
Received for publication January 30, 2002.
Accepted for publication August 27, 2002.
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