Phosphorylation of Threonine 333 Regulates Trafficking of the Human sst₅ Somatostatin Receptor

Abstract

The frequent overexpression of the somatostatin receptors sst₂ and sst₅ in neuroendocrine tumors provides the molecular basis for therapeutic application of novel multireceptor somatostatin analogs. Although the phosphorylation of the carboxyl-terminal region of the sst₂ receptor has been studied in detail, little is known about the agonist-induced regulation of the human sst₅ receptor. Here, we have generated phosphosite-specific antibodies for the carboxyl-terminal threonines 333 (T333) and 347 (T347), which enabled us to selectively detect either the T333-phosphorylated or the T347-phosphorylated form of sst₅. We show that agonist-mediated phosphorylation occurs at T333, whereas T347 is constitutively phosphorylated in the absence of agonist. We further demonstrate that the multireceptor somatostatin analog pasireotide and the sst₅-selective ligand L-817,818 but not octreotide or KE108 were able to promote a detectable T333 phosphorylation. Interestingly, BIM-23268 was the only sst₅ agonist that was able to stimulate T333 phosphorylation to the same extent as natural somatostatin. Agonist-induced T333 phosphorylation was dose-dependent and selectively mediated by G protein-coupled receptor kinase 2. Similar to that observed for the sst₂ receptor, phosphorylation of sst₅ occurred within seconds. However, unlike that seen for the sst₂ receptor, dephosphorylation and recycling of sst₅ were rapidly completed within minutes. We also identify protein phosphatase 1γ as G protein-coupled receptor phosphatase for the sst₅ receptor. Together, we provide direct evidence for agonist-selective phosphorylation of carboxyl-terminal T333. In addition, we identify G protein-coupled receptor kinase 2-mediated phosphorylation and protein phosphatase 1γ-mediated dephosphorylation of T333 as key regulators of rapid internalization and recycling of the human sst₅ receptor.

Somatostatin (SS-14) is a cyclic peptide that regulates an array of physiologic functions via inhibition of secretion of hormones such as GH, TSH, ACTH, insulin, and glucagon (1). SS-14 is the natural ligand of a family of 5 G protein-coupled receptors named sst₁–sst₅ (2). Given its short half-life in human plasma, metabolically stable somatostatin analogs have been developed. Among these, octreotide and lanreotide predominantly mediate their effects via the sst₂ receptor. In clinical practice, octreotide and lanreotide are used as first-choice medical treatment of neuroendocrine tumors such as GH-secreting adenomas and carcinoid (3). Loss of octreotide response in these tumors occurs because of diminished expression of sst₂, whereas sst₅ expression persists (4). Recently, the novel multireceptor somatostatin analog, pasireotide (SOM230), has been synthesized (5). In contrast to octreotide, pasireotide exhibits particularly high subnanomolar affinity to sst₅ (6). Pasireotide has recently been approved for the treatment of Cushing's disease, a condition with known sst₅ overexpression (7). Pasireotide is also under clinical evaluation for the treatment of acromegaly and octreotide-resistant carcinoid tumors (8, 9).

We have recently used phosphosite-specific antibodies to examine agonist-induced phosphorylation of the sst₂ receptor. We found that SS-14 promotes the phosphorylation of at least 6 carboxyl-terminal serine and threonine residues, namely, S341, S343, T353, T354, T356, and T359 (10–12). This phosphorylation is mediated by G protein-coupled receptor kinase 2 (GRK2) and GRK3 and followed by rapid cointernalization of the receptor and β-arrestin into the same endocytic vesicles (12, 13). Dephosphorylation of sst₂ is initiated directly after receptor activation at or near the plasma membrane and is mediated by protein phosphatase 1β (PP1β) (14). Although we have recently provided evidence for phosphorylation of threonine 333 (T333) (10), our knowledge about the functional role of carboxyl-terminal phosphorylation of the sst₅ receptor is limited. In fact, contrasting findings have been reported regarding the role of the carboxyl-terminus in sst₅ internalization (15, 16). Although truncation of the carboxyl-terminal tail to 318, 328, and 338 residues has been observed to inhibit receptor internalization in Chinese hamster ovary K1 cells (15), the same truncations resulted in a progressive increase in sst₅ internalization in rat pituitary GH3 cells (16). In the present study, we have examined the primary structure of the sst₅ carboxyl-terminal tail. A comparison to the sst₂ receptor revealed the presence of 2 potential phosphorylation sites, namely T333 and threonine 347 (T347), in the region that corresponds to the phosphorylation-sensitive domain of the sst₂ receptor. Consequently, we have generated phosphosite-specific antibodies, which enabled us to provide direct evidence for carboxyl-terminal phosphorylation of the sst₅ receptor. In addition, we identify kinases and phosphatases involved in the regulation of agonist-dependent phosphorylation of the sst₅ receptor.

Materials and Methods

Antibodies and reagents

Phosphosite-specific antibodies for the T333-phosphorylated form of sst₅ were generated against the following sequence that contained a phosphorylated threonine residue: KDATA(pT)EPRPD. This sequence corresponds to 328–338 of the human sst₅. Phosphosite-specific antibodies for the T347-phosphorylated form of sst₅ were generated against the following sequence that contained a phosphorylated threonine residue: QQEA(pT)PPAHR. This sequence corresponds to 343–352 of the human sst₅. The peptides were purified by HPLC and coupled to keyhole limpet hemocyanin via a carboxyl-terminally added cystein residue. The conjugates were mixed 1:1 with Freund's adjuvant and injected into 1 group of 4 rabbits {3567–3570} for anti-pT333 antibody production and 1 group of 3 rabbits for anti-pT347 {3563–3565}. Animals were injected at 4-week intervals, and serum was obtained 2 weeks after immunizations beginning with the second injection. The specificity of the antisera was initially tested using dot-blot analysis. For subsequent analysis, antibodies were affinity purified against their immunizing peptide as well as against the nonphosphorylated peptide using the SulfoLink kit (Thermo Scientific, Rockford, Illinois). Equal loading of the gels was confirmed using the phosphorylation-independent rabbit monoclonal anti-sst5 antibody {UMB-4}, which was extensively characterized previously (17). The phosphorylation-independent rabbit monoclonal anti-sst₂ antibody {UMB-1} and the phosphosite-specific sst_2A antibodies anti-pS341/pS343 {3157}, anti-pT353/pT354 {0521}, anti-pT356/pT359 {0522}, and the rabbit polyclonal anti-hemagglutinin (HA) antibody were generated and extensively characterized as previously described (10–12, 18).

Anti-GRK2 (sc-562), anti-GRK3 (sc-563), anti-GRK5 (sc-565), anti-GRK6 (sc-566), anti-PP1α (sc-6104), anti-PP1β (sc-6106), and anti-PP1γ (sc-6108) antibodies were obtained from Santa Cruz Biotechnology (Heidelberg, Germany). SS-14, phorbol 12-myristate 13-acetate (PMA), and forskolin were obtained from Sigma-Aldrich (St Louis, Missouri). Octreotide and pasireotide were kind gifts from Novartis (Basel, Switzerland). L-817,818, a selective ligand for sst₅, was provided by Merck (Whitehouse Station, New Jersey). BIM-23268, BIM-23A760, BIM-23627, and BIM-23454 were provided by Ipsen (Milford, Massachusetts). KE108 was obtained from Bachem (Bubendorf, Switzerland).

Cell culture and transfection

Human embryonic kidney (HEK) 293 cells were obtained from the German Resource Centre for Biological Material (DSMZ, Braunschweig, Germany). HEK 293 cells were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum. Cells were transfected with a plasmid encoding for an HA-tagged human sst₅ receptor (cDNA Resource Center, Rolla, Montana) using LipofectAMINE 2000 according to the instructions of the manufacturer (Invitrogen, Carlsbad, California). Stable transfectants were selected in the presence of 400 μg/mL G418. HEK 293 cells stably expressing human sst₅ receptor were characterized using radioligand-binding assays, cAMP assays, Western blot analysis, and immunocytochemistry as described previously (13, 19). HA-tagged T333A and T333A/T347A mutants of the human sst₅ receptor were generated by gene synthesis and obtained from imaGenes (Berlin, Germany).

Western blot analysis

Cells were seeded onto 60-mm dishes and grown to 80% confluence. After the indicated treatment with SS-14, L-817,818, octreotide, pasireotide, KE108, PMA, or forskolin, cells were lysed in detergent buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 5 mM EDTA, 10 mM NaF, 10 mM disodium pyrophosphate, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS) in the presence of protease and phosphatase inhibitors, Complete mini, and PhosSTOP (Roche Diagnostics, Mannheim, Germany). When indicated, cells were incubated with 5 nM, 10 nM, or 30 nM calyculin A or with 5 nM, 50 nM, or 500 nM okadaic acid for 10 minutes before agonist exposure. All phosphorylation and dephosphorylation assays were performed at both physiologic temperature (37°C) and at room temperature (22°C) for the indicated time periods. Glycosylated proteins were partially enriched using wheat germ lectin-agarose beads as described (20, 21). Proteins were eluted from the beads using SDS-sample buffer for 20 minutes at 47°C. When indicated, samples were dephosphorylated with lambda protein phosphatase (New England Biolabs, Frankfurt, Germany) for 2 hours at 37°C. Samples were resolved on 7.5% SDS-polyacrylamide gels, and after electroblotting, membranes were incubated with either 0.1 μg/mL anti-pT333 {3567} or 0.1 μg/mL anti-pT347 {3564} followed by detection using an enhanced chemiluminescence detection system (Amersham, Braunschweig, Germany). Blots were subsequently stripped and reprobed with anti-sst₅ antibody {UMB-4} to confirm equal loading of the gels. When indicated, cells were washed 3 times with 2 mL ice-cold PBS (Washout) to remove SS-14. Protein bands on Western blots were captured and quantified using the Fusion FX7 chemiluminscence reader (Peqlab, Erlangen, Germany). Blots were subsequently exposed to x-ray film. Films exposed in the linear range were then densitized using ImageJ 1.45. Data were analyzed using GraphPad Prism 4.0 software (La Jolla, California).

Immunocytochemistry

Cells were grown on poly-l-lysine-coated coverslips overnight. After drug treatment, cells were fixed with 4% paraformaldehyde and 0.2% picric acid in phosphate buffer (pH 6.9) for 30 minutes at room temperature and washed several times. Cells were permeabilized and then incubated with anti-sst₅ antibody {UMB-4} followed by Alexa488-conjugated secondary antibodies (Amersham). Specimens were mounted and examined using a Zeiss LSM510 META laser scanning confocal microscope (Zeiss, Jena, Germany).

Quantitative internalization assays

Cells were seeded onto 24-well plates. On the next day, cells were preincubated with anti-HA antibody for 2 hours at 4°C. Cells were then transferred to 37°C, exposed to agonist, fixed, and developed with peroxidase-conjugated secondary antibody as described previously (12, 22).

Small interfering RNA silencing of gene expression

Chemically synthesized double-stranded small interfering RNA (siRNA) duplexes (with 3′ dTdT overhangs) were purchased from Qiagen (Hilden, Germany) for the following targets: GRK2 (5′-CCGGGAGATCTTCGACTCATA-3′ and 5′-AAGAAGTACGAGAAGCTGGAG-3′), GRK3 (5′-AAGCAAGCTGTAGAACACGTA-3′ and 5′-GCAGAAGTCGACAAATTTA-3′), GRK5 (5′-AGCGTCATAACTAGAACTGAA-3′ and 5′-AAGCCGTGCAAAGAACTCTTT-3′), GRK6 (5′-AACACCTTCAGGCAATACCGA-3′ and 5′-AACAGTAGGTTTGTAGTGAGC-3′), PP1α catalytic subunit (5′-AAGAGACGCTACAACATCAAA-3′), PP1β catalytic subunit (5′-TACGAGGATGTCGTCCAGGAA-3′ and 5′-GTTCGAGGCTTATGTATCA-3′), PP1γ catalytic subunit (5′-AACATCGACAGCATTATCCAA-3′ and 5′-AGAGGCAGTTGGTCACTCT-3′), and a nonsilencing RNA duplex (5′-GCTTAGGAGCATTAGTAAA-3′ or 5′-AAA CTC TAT CTG CAC GCT GAC-3′). HEK 293 cells were transfected with 150 nM siRNA for single transfection or with 100 nM of each siRNA for double transfection using HiPerFect (Qiagen). Silencing was quantified by immunoblotting. All experiments showed protein levels reduced by ≥80%.

GTPγS binding assays

Cells were harvested and lysed as described above except that a lysis buffer containing 50 mM Tris, 10 mM EDTA, and 1 mM EGTA (pH 7.4) was used (10). The resulting pellet was resuspended in assay buffer (20 mM HEPES, 100 mM NaCl, 10 mM MgCl₂, pH 7.4). Aliquots containing 30 μg protein were incubated with 3 μM GDP and 0.05 nM [³⁵S]GTPγS (specific activity, 43.3 TBq/mmol; Perkin-Elmer, Waltham, Massachusetts) in the presence or absence of either SS-14, octreotide, pasireotide, L-817,818, or KE108 in concentrations ranging from 10⁻¹² to 10⁻⁶ M. Assays were carried out in a final volume of 1 mL for 30 minutes at 30°C under continuous agitation. Nonspecific binding was determined in the presence of 10 μM unlabeled GTPγS. The incubation was terminated by the addition of ice-cold buffer and rapid vacuum filtration through glass fiber filters. Filters were rinsed twice with washing buffer (50 mM Tris-HCl, pH 7.4) and dried. A scintillation mixture was added, and radioactivity was determined using a β-counter (1600 TR; Packard, Milan, Italy).

Data analysis

Data were analyzed using GraphPad Prism 4.0 software. Statistical analysis was carried out with 2-way ANOVA followed by the Bonferroni posttest. P values of <.05 were considered statistically significant.

Results

We have recently shown that agonist-induced phosphorylation of sst₂ occurs at a specific cluster of serine and threonine residues within the carboxyl-terminal tail of the receptor (12, 13). We hypothesized that this region based on its distance from the seventh transmembrane domain would be particularly accessible for GRK-mediated phosphorylation. Therefore, we have examined the primary structures of sst₂ and sst₅ (Figure 1, A and B). We found only 2 potential phosphate acceptor sites, namely T333 and T347, in the region that corresponds to the phosphorylation-sensitive domain of the sst₂ receptor. Consequently, we generated phosphosite-specific antibodies to pT333 and pT347. These antisera were affinity-purified against their immunizing peptides and initially tested in dot-blot assays using the phosphopeptides and the corresponding nonphosphopeptides (not shown). All antibodies, which clearly detected their respective phosphopeptide and did not cross-react with the corresponding nonphosphopeptide, were further characterized in Western blot assays using HEK 293 cells stably expressing the sst₅ receptor. In Western blots from these cells, the sst₅ receptor is detected as a broad band migrating at molecular weight of 55 000–70 000 (4, 17, 21). This band represents the N-glycosylated form of sst₅. After enzymatic deglycosylation using PNGase F, the sst₅ receptor is detected at an approximate molecular weight of 40 000, which corresponds well to the expected size of the unglycosylated sst₅ receptor (17).

Figure 1.

Characterization of phosphosite-specific sst₅ antibodies[b]. A and B, Schematic representation of the carboxyl-terminal region of human sst₂ and sst₅ receptors. Phosphate acceptor sites targeted for the generation of phosphosite-specific antibodies are marked in gray. C, HEK 293 cells stably expressing the sst₅ receptor were either not exposed (−) or exposed (+) to 1 μM SS-14 for 30 minutes. Cells were lysed and immunoblotted with the anti-pT333 antibodies {3567}, {3568}, and {3569} or anti-pT347 antibodies {3563}, {3564}, and {3565} at a concentration of 0.1 μg/mL (upper panel). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} at a dilution of 1:500 to confirm equal loading of the gels (lower panel). Note that T347 phosphorylation is detectable in both SS-14-treated and untreated cells. D, HEK 293 cells stably expressing the sst₅ receptor were either not exposed (−) or exposed (+) to 1 μM SS-14. Cells were lysed and proteins were either dephosphorylated (+) using lambda protein phosphatase or not dephosphorylated (−). Samples were then immunoblotted with the anti-pT333 antibody {3567} or the anti-pT347 antibody {3564} (upper panel). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} to confirm equal loading of the gels (lower panel). E, HEK 293 cells stably expressing the wild-type sst₅ receptor (sst5-WT) or the T333A/T347A mutant of sst₅ (T333A/T347A) were either not exposed (−) or exposed (+) to 1 μM SS-14. Samples were then immunoblotted with the anti-pT333 antibody {3567} or the anti-pT347 antibody {3564} (upper panel). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} (lower panel). Shown are representative results from 1 of 3 independent experiments. The position of molecular mass markers is indicated on the left (in kilodaltons).

As depicted in Figure 1C, the anti-pT333 antibodies {3567}, {3568}, and {3569} detected the T333-phosphorylated form of sst₅ only in SS-14-treated cells but not in untreated cells. The signal of anti-pT333 antibody {3567} was very robust without detectable background. Consequently, the anti-pT333 antibody {3567} was used throughout this study. The anti-pT347 antibodies {3563}, {3564}, and {3565} detected the T347-phosphorylated form of the receptor in both SS-14-treated and untreated cells. All of them showed a robust signal without detectable background (Figure 1C). To test whether the band detected in untreated cells by the anti-pT347 antibody {3564} would represent the T347-phosphorylated form of sst₅, we used lambda phosphatase to dephosphorylate the receptors. As depicted in Figure 1D, after phosphatase treatment, both the anti-pT333 antibody {3567} and the anti-pT347 antibody {3564} were no longer able to detect their cognate forms of phosphorylated sst₅ receptors, whereas the receptor protein was still detectable using the phosphorylation state-independent antibody {UMB-4}. Moreover, when T333 and T347 were mutated to alanine, the anti-pT333 antibody {3567} as well as the anti-pT347 antibody {3564} were no longer able to detect phosphorylated sst₅ receptors (Figure 1E). These findings strongly suggest that T347 is constitutively phosphorylated in the absence of agonist. In contrast, T333 is involved in agonist-induced phosphorylation.

To examine role of T333 and T347 phosphorylation in sst₅ receptor sequestration, we generated T333A and T333A/T347A mutant receptors. These sst₅ mutants were expressed and SS-14-stimulated receptor internalization was monitored at 37°C using confocal microscopy and a quantitative surface ELISA. The results shown in Figure 2 reveal that mutation of T333 to alanine was sufficient to block agonist-driven internalization, suggesting that T333 phosphorylation is a key regulatory event during sst₅ receptor sequestration.

Figure 2.

Phosphorylation of T333 is required for agonist-induced sst₅ internalization. A, HEK 293 cells stably expressing the wild-type sst₅ receptor (sst5-WT), the T333A-mutant (T333A), or the T333A/T347A mutant of sst₅ (T333A/T347A) were either not exposed (−) or exposed to 1 μM SS-14 for 30 minutes. Cells were fixed and stained with the phosphorylation-independent anti-sst₅ antibody {UMB-4}. Shown are representative results from 1 of 3 independent experiments. Scale bar, 20 μm. B, HEK 293 cells stably expressing the wild-type sst₅ receptor (sst5-WT), the T333A-mutant (T333A), or the T333A/T347A mutant of sst₅ (T333A/T347A) were either not exposed (−) or exposed to 1 μM SS-14 for 30 minutes. Internalization of the different sst₅ receptors was determined by ELISA. Results were expressed as percentage internalization of surface receptors. Results were analyzed by 2-way ANOVA followed by the Bonferroni posttest (*P < .05).

We then assessed whether other sst₅ agonists would be able to induce T333 phosphorylation. We found that the multireceptor somatostatin analog pasireotide and the sst₅-selective ligand L-817,818 were able to promote a clearly detectable T333 phosphorylation (Figure 3, left panel). In contrast, no such signal was seen after incubation with octreotide or KE108. However, none of the compounds tested was able to stimulate T333 phosphorylation of sst₅ to the same extent as the natural ligand SS-14 (Figure 3, left panel). Interestingly, the only compound that was able to stimulate T333 phosphorylation to a similar degree as SS-14 was the sst₅-selective agonist BIM-23268 (Figure 3, second panel). In general, there was an excellent correlation between T333 phosphorylation and sst₅ receptor internalization (Figure 3, left panel). Nevertheless, all compounds were able to stimulate GTPγS binding under otherwise identical conditions (Figure 3, left panel). This suggests that many currently available sst₅ agonists exhibit a bias toward G protein signaling. We further examined dose-dependent T333 phosphorylation of sst₅ after exposure to various concentrations of SS-14 ranging from 10⁻⁹ to 10⁻⁵ M. As depicted in Figure 3 (third panel), a weak phosphorylation signal was first detectable at 10⁻⁸ M SS-14. T333 phosphorylation reached a maximum between 10⁻⁷ and 10⁻⁶ M SS-14 (Figure 3, third panel). We have previously reported that for the human sst₅ receptor maximal inhibition of cAMP accumulation is already observed at 10⁻⁸ M SS-14 (22). The higher activity of SS-14 to stimulate sst₅-dependent signaling reflects a high receptor reserve in this experimental system. Consequently, maximal stimulation of sst₅ phosphorylation and internalization requires approximately 10-fold higher concentrations of SS-14 (12, 22).

Figure 3.

Agonist-selective T333 phosphorylation of sst₅. Left panel, HEK 293 cells stably expressing the sst₅ receptor were either not exposed (−) or exposed to 1 μM SS-14, 1 μM L-817,818, 1 μM octreotide, 1 μM pasireotide, or 1 μM KE108 for 5 minutes at room temperature (22°C). G protein signaling was determined by a GTPγS binding assay. Results were quantified and expressed as percentage of maximal G protein binding in unstimulated cells, which was set at 100%. Internalization was determined by surface ELISA. Results were expressed as percentage internalization of surface receptors. Second panel, HEK 293 cells stably expressing the sst₅ receptor were either not exposed (−) or exposed to 1 μM SS-14, 1 μM BIM-23A760, or 1 μM BIM-23268 for 5 minutes at room temperature (22°C). Third panel, HEK 293 cells stably expressing the sst₅ receptor were either not exposed (−) or exposed to the indicated concentrations of SS-14 for 5 minutes at room temperature. Fourth panel, HEK 293 cells stably expressing the sst₅ receptor were either not exposed or exposed to 1 μM SS-14, 0.1 μM PMA, or 10 μM forskolin for 20 minutes at room temperature. Right panel, HEK 293 cells stably expressing the sst₅ receptor were either not exposed or exposed to 1 μM SS-14, 1 μM SS-14 plus 10 μM BIM-23627, or 1 μM SS-14 plus 10 μM BIM-23454 for 10 minutes. Cells were lysed and immunoblotted with the anti-pT333 antibodies {3567} (pT333). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} to confirm equal loading of the gel (sst5). Representative results from 1 of 3 independent experiments are shown. The position of molecular mass markers is indicated on the left (in kilodaltons).

Phosphorylation of G protein-coupled receptor (GPCRs) can occur via specific GRKs or second messenger-activated kinases (eg, protein kinase A or protein kinase C). We therefore treated sst₅-expressing HEK 293 cells with forskolin or PMA and examined T333 phosphorylation. We found that neither 100 nM PMA nor 10 μM forskolin produced a detectable phosphorylation of T333 (Figure 3, fourth panel). We also tested whether SS-14-induced T333 phosphorylation could be inhibited by application of BIM-23627 or BIM-23454, 2 sst₂ antagonists known to bind to the sst₅ receptor as well. When cells were coincubated with SS-14 and either antagonist for 10 minutes, a clear reduction in T333 phosphorylation was noted (Figure 3, right panel). It should be noted that the inhibition of T333 phosphorylation observed with BIM-23627 was somewhat stronger than for BIM-23454, which may simply reflect its higher affinity to the sst₅ receptor. Nevertheless, none of these compounds was able to block T333 phosphorylation completely (Figure 3, right panel).

We then elucidated the fundamental question, which kinases are involved in T333 phosphorylation. We used siRNA to knockdown gene expression of GRK2, GRK3, GRK5, and GRK6. We have recently observed that in promoting agonist-induced sst₂ phosphorylation GRK2 and GKR3 can function as a redundant phosphorylation system (12). Consequently, we also tested combined siRNA knockdown of GRK2 plus GRK3 and GRK5 plus GRK6. Interestingly, knockdown of GRK2 but not of GRK3 resulted in a robust inhibition of SS-14-induced T333 phosphorylation (Figure 4A). Quantitative analysis revealed that T333 phosphorylation was reduced by ∼40% after knockdown of GRK2 and by ∼50% after knockdown of both GRK2 and GRK3 (Figure 4A). No such inhibition was observed after knockdown of GRK5 or GRK6 (Figure 4B).

Figure 4.

GRK2 is responsible for agonist-induced T333 phosphorylation. A, HEK 293 cells stably expressing the sst₅ receptor were transfected with siRNA targeted to GRK2, GRK3, GRK2, and GRK3 or nonsilencing siRNA control (SCR) for 72 hours and then exposed to 1 μM SS-14 for 10 minutes. B, HEK 293 cells stably expressing the sst₅ receptor were transfected with siRNA targeted to GRK5, GRK6, GRK5, and GRK6 or nonsilencing siRNA control (SCR) for 72 hours and then exposed to 1 μM SS-14 for 10 minutes. Cells were lysed and immunoblotted with the anti-pT333 antibodies {3567} (pT333). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} to confirm equal loading of the gels (sst5). siRNA knock-down of GRK2, GRK3, GRK5, and GRK6 was confirmed by Western blot. Blots were quantified and expressed as percentage of maximal phosphorylation in SCR-transfected cells. Data correspond to the mean ± SEM from 3 independent experiments. Results were analyzed by 2-way ANOVA followed by the Bonferroni posttest (*P < .05). siRNA knock-down of GRK2 was confirmed by Western blot. The positions of molecular mass markers are indicated on the left (in kilodaltons).

Among the 5 somatostatin receptor subtypes, only sst₂ and sst₅ are proven drug targets for stable somatostatin analogs. We have recently compared the signaling and trafficking properties of human somatostatin receptors (22). In the presence of their endogenous ligand, sst₂ and sst₅ receptors exhibited very similar abilities to activate ERK1/2 and to inhibit forskolin-stimulated cAMP formation (22). Under identical conditions, sst₂ receptors exhibited a nearly complete internalization, whereas sst₅ receptors were only partially internalized (22). We therefore compared the patterns of sst₅ and sst₂ trafficking with the temporal dynamics of their phosphorylation and dephosphorylation. First, we performed an immunocytochemical analysis of receptor internalization and recycling at 37°C. Confocal microscopic images depicted in Figure 5 show that sst₅ internalization was clearly detectable. However, after 15 minutes and 30 minutes, only a minor proportion of sst₅ receptors (∼20%–30%) was removed from the plasma membrane and internalized into endocytic vesicles (Figures 3 and 5). After washout of the ligand, sst₅ receptors were able to complete their recycling to the plasma membrane within 30 minutes (Figure 5). Although nearly all sst₂ receptors were internalized within 15 minutes, sst₂ receptors were not able to complete their trafficking to the plasma membrane within 60 minutes in the absence of agonist (Figure 5).

Figure 5.

Comparison of sst₅ and sst₂ receptor internalization and recycling. HEK 293 cells stably expressing the sst₅ or the sst₂ receptor were exposed to 1 μM SS-14 for 0, 15, or 30 minutes at physiologic temperature (37°C). Cells were subsequently washed 3 times with cold PBS (Washout) and incubated in the absence of agonist for 30 or 60 minutes at physiologic temperature (37°C). Cells were fixed and stained immunofluorescently and examined by confocal microscopy. Representative results from 1 of 3 independent experiments are shown. Scale bar, 20 μm.

We then used a quantitative surface ELISA assay to resolve the time courses of sst₅ and sst₂ receptor internalization at physiologic temperature (37°C). As depicted in Figure 6, for both receptors internalization reached a maximum between 15 minutes and 30 minutes. For sst₅ no more than 30% to 40% of surface receptors were removed from the plasma membrane (Figure 6A). In contrast, maximal internalization of sst₂ receptors was between 80% and 90% (Figure 6B). All phosphorylation and dephosphorylation assays were performed at both physiologic temperature (37°C) and at room temperature (22°C). Initial experiments revealed that phosphorylation of both receptors occurred at physiologic temperature (37°C) much more rapidly than receptor internalization. To resolve differences in the phosphorylation of individual sites of both receptors in more detail, we decreased the temperature to room temperature (22°C). In the presence of SS-14, T333 phosphorylation of sst₅ was detectable already after 20 seconds and reached a maximum after 1 to 2 minutes (Figure 6A). Similar to that observed at physiologic temperature, constitutive phosphorylation of T347 was also detectable at room temperature in the absence of agonist. Within minutes of agonist exposure, we observed a small increase in T347 phosphorylation, suggesting that T347 can also be a substrate for homologous agonist-driven phosphorylation (Figure 6A). Similar to that observed for the sst₅ receptor, robust agonist-induced phosphorylation of sst₂ was detectable within 20 seconds (Figure 6B). Unlike sst₅, phosphorylation of the sst₂ receptor occurred within 20 seconds on all known phosphate acceptor sites, including S341/S343, T353/T354, and T356/T359. When sst₅-expressing cells were exposed to SS-14 for 5 minutes, washed, and then incubated in agonist-free medium, T333 dephosphorylation occurred very rapidly (Figure 7A). In contrast, T347 phosphorylation was to a lesser extent still detectable even after prolonged incubation (60 minutes) in the absence of agonist (Figure 7A). Interestingly, dephosphorylation of individual sst₂ phosphate acceptor sites occurred with distinct temporal dynamics. Although T353/T354 dephosphorylation occurred rapidly, T356/T359 dephosphorylation was delayed and S341/S343 dephosphorylation was not detectable for at least 2 hours after SS-14 washout (Figure 7B). These results indicate that sst₅ and sst₂ exhibit strikingly different patterns of dephosphorylation and recycling. Although the fast sst₅ trafficking correlates with the rapid T333 phosphorylation and dephosphorylation, the sst₂ recycling appears to be delayed due to its slow dephosphorylation.

Figure 6.

Comparison of agonist-induced phosphorylation of sst₅ and sst₂. A, HEK 293 cells stably expressing the sst₅ receptor were exposed to 1 μM SS-14 at 37°C for the indicated time periods. Internalization of sst₅ was determined by ELISA. Results were expressed as percentage internalization of surface receptors (top). HEK 293 cells stably expressing the sst₅ receptor were exposed to 1 μM SS-14 at room temperature for the indicated time periods. Cells were lysed and immunoblotted with the anti-pT333 antibody {3567} (pT333) or the anti-pT347 antibody {3564} (pT347). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} to confirm equal loading of the gels (sst5). B, HEK 293 cells stably expressing the sst₂ receptor were exposed to 1 μM SS-14 at 37°C for the indicated time periods. Internalization of sst₂ was determined by ELISA. Results were expressed as percentage internalization of surface receptors (top). HEK 293 cells stably expressing the sst₂ receptor were exposed to 1 μM SS-14 at room temperature for the indicated time periods. Cells were lysed and immunoblotted with the anti-pS341/S343 antibody {3157} (pS341/S343) or the anti-pT353/T354 antibody {0521} (pT353/T354) or the anti-pT356/T359 antibody {0522} (pT356/T359). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₂ antibody {UMB-1} to confirm equal loading of the gels (sst2). Shown are representative results from 1 of 3 independent experiments. The position of molecular mass markers is indicated on the left (in kilodaltons).

Figure 7.

Comparison of sst₅ and sst₂ dephosphorylation. A, HEK 293 cells stably expressing the sst₅ receptor were exposed to 1 μM SS-14 for 5 minutes, washed, and incubated in the absence of agonist for 0, 1, 2, 5, 10, 15, 30, 45, or 60 minutes. Cells were lysed and immunoblotted with the anti-pT333 antibody {3567} (pT333) and the anti-pT347 antibody {3564} (pT347). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₅ antibody {UMB-4} to confirm equal loading of the gels (sst5). B, HEK 293 cells stably expressing the sst₂ receptor were exposed to 1 μM SS-14 for 5 minutes, washed, and incubated in the absence of agonist for 0, 2, 5, 10, 20, 40, 60, or 120 minutes. Cells were lysed and immunoblotted with the anti-pS341/S343 antibody {3157} (pS341/S343) or the anti-pT353/T354 antibody {0521} (pT353/T354) or the anti-pT356/T359 antibody {0522} (pT356/T359). Blots were stripped and reprobed with the phosphorylation-independent anti-sst₂ antibody {UMB-1} to confirm equal loading of the gels (sst2). Shown are representative results from 1 of 3 independent experiments. The position of molecular mass markers is indicated on the left (in kilodaltons).

Recently, we identified PP1β as GPCR phosphatase for the sst₂ using a combination of chemical inhibitors and siRNA knockdown screening (14). Consequently, we used the same approach to identify the GPCR phosphatase responsible for rapid T333 dephosphorylation of sst₅. When stably transfected HEK 293 cells were exposed to increasing concentrations of protein phosphatase inhibitors, sst₅ dephosphorylation was inhibited in a dose-dependent manner only by calyculin A but not by okadaic acid (Figure 8). Both calyculin A and okadaic acid can effectively block the protein phosphatases PP2, PP4, and PP5 (23). In contrast to okadaic acid, calyculin A is also a potent inhibitor of PP1 activity (23). Thus, the present data suggest that PP1 activity is required for rapid T333 dephosphorylation. To date, 3 distinct catalytic subunits, α, β, and γ, are known for PP1. To elucidate which of these PP1 isoforms is involved in sst₅ dephosphorylation, we performed siRNA knockdown experiments. As depicted in Figure 9, only PP1γ knockdown resulted in a robust inhibition of sst₅ dephosphorylation. In contrast, transfection of PP1α or PP1β siRNA had no effect on sst₅ dephosphorylation (Figure 9). These results indicate that PP1γ is the GPCR phosphatase responsible for rapid T333 dephosphorylation of sst₅.

Figure 8.

Calyculin A but not okadaic acid prevents sst₅ dephosphorylation. HEK 293 cells stably expressing sst₅ were treated with calyculin A (A) or okadaic acid (B) for 10 minutes at the indicated concentrations and then exposed to 1 μM SS-14 for 5 minutes in the presence of calyculin A or okadaic acid. Cells were washed 3 times and then incubated for 0, 2, or 7 minutes in the presence of calyculin A or okadaic acid. The levels of T333-phosphorylated sst₅ receptors (pT333) and total sst₅ receptors (sst5) were then determined by Western blot analysis. Blots were quantified and expressed as percentage of maximal phosphorylation in untreated cells, which was set at 100%. Data correspond to the mean ± SEM from 4 independent experiments. Note that calyculin A but not okadaic acid inhibited sst₅ receptor dephosphorylation in a dose-dependent manner. Western blots shown are representative of 4 independent experiments for each condition. The positions of molecular mass markers are indicated on the left (kilodaltons).

Figure 9.

PP1γ catalyzes pT333 dephosphorylation. A, HEK 293 cells stably expressing the sst₅ receptor were transfected with siRNA targeted to PP1α, PP1β, PP1γ, or nonsilencing siRNA control (SCR) for 72 hours and then exposed to 1 μM SS-14 for 5 minutes. Cells were washed 3 times and then incubated for 0, 2, or 5 minutes in the absence of agonist. Cells were lysed and immunoblotted with anti-pT333 antibodies (pT333). Blots were stripped and reprobed with the phosphorylation-independent anti-sst5 antibody {UMB-4} to confirm equal loading of the gels (sst5). T333 phosphorylation was quantified and expressed as percentage of maximal phosphorylation in SCR-transfected cells, which was set at 100%. Data correspond to the mean ± SEM from 5 independent experiments. Results were analyzed by 2-way ANOVA followed by the Bonferroni posttest (*P < .05). Note that transfection with PP1γ siRNA resulted in a significant inhibition of T333 dephosphorylation. Western blots shown are representative of 5 independent experiments. B, siRNA knockdown of PP1 was confirmed by Western blot using subunit-specific PP1 antibodies. The positions of molecular mass markers are indicated on the left (in kilodaltons).

Discussion

The human sst₅ receptor is a major drug target for the novel multireceptor somatostatin analog pasireotide. Compared to the closely related sst₂ receptor, however, little is known about its agonist-driven regulation. In the present study, we generated and extensively characterized phosphosite-specific antibodies to T333 and T347, the only 2 potential phosphate acceptor sites located in the region that corresponds to the sst₂ phosphorylation motif. We show that T333 is rapidly phosphorylated and dephosphorylated in an agonist-dependent manner. We also show that mutation of T333 strongly reduced sst₅ internalization. Interestingly, mutation of phosphate acceptor sites within the third intracellular loop including S242 and T247 has also been shown to partially inhibit receptor internalization (16). Thus, it is possible that these sites play a complementary role in regulating sst₅ internalization.

We also found that T347 is constitutively phosphorylated in the absence of agonist. We identify GRK2 as the kinase responsible for T333 phosphorylation and PP1γ as the phosphatase responsible for T333 dephosphorylation. For many GPCRs carboxyl-terminal phosphorylation is crucial for their interaction with β-arrestin and subsequent receptor sequestration (24). In fact, for sst₅ there is an excellent correlation between extent and temporal dynamics of carboxyl-terminal T333 phosphorylation and its trafficking properties. After agonist exposure, sst₅ is phosphorylated and β-arrestin is recruited to the receptor (13). T333 becomes rapidly dephosphorylated, leading to disruption of the β-arrestin-receptor complex, thus permitting accelerated recycling of the receptor to the plasma membrane (13).

The present study also uncovers striking differences in the phosphorylation and trafficking patterns of sst₅ and sst₂ receptors. In contrast to sst₅, agonist-induced phosphorylation of sst₂ occurs on 6 carboxyl-terminal phosphate acceptor sites; β-arrestin forms stable complexes with the sst₂ receptor, which in turn internalize together into the same endocytic vesicles (12, 13). Consequently, dephosphorylation and recycling of sst₂ occur at a much slower rate than that observed for sst₅. Although dephosphorylation of the sst₅ receptor and of the μ-opioid receptor involves PP1γ (25, 26), dephosphorylation of sst₂ requires PP1β activity (14). This is an intriguing finding. At present, very little is known about the molecular mechanisms of GPCR dephosphorylation. After our initial observation of PP1β as GPCR phosphatase for sst₂, PP1γ is the second GPCR phosphatase identified (14, 26). However, it is unclear which mechanisms regulate phosphatase specificity. It is possible that either carboxyl-terminal phosphorylation motifs, specific sequences within the intracellular loops of the receptor or the β-arrestin trafficking patterns, may contribute to phosphatase selection.

We have recently generated the novel rabbit monoclonal antibodies UMB-1 and UMB-4 directed to human sst₂ and human sst₅, respectively (17, 18). The use of UMB-1 and UMB-4 permitted us to gain novel insights into sst₂ and sst₅ receptor expression and function. We found that the sst₅ receptor is present on the plasma membrane of all GH-producing cells as well as all ACTH-producing cells in the anterior pituitary, whereas sst₂ was only present on a subpopulation of GH-producing cells (17). Both sst₅ and sst₂ were detected on the plasma membrane of all insulin-producing cells as well as all glucagon-producing cells in human pancreatic islets (17). We also observed a high prevalence of both sst₅ and sst₂ receptors in pituitary adenomas and neuroendocrine tumors (17). Octreotide and lanreotide are first-line agents in the medical management of clinical conditions with predominant sst₂ expression such as GH-secreting adenomas and carcinoid tumors (3, 27). Pasireotide is effective in conditions with known sst₅ overexpression including Cushing's disease, acromegaly, and octreotide-resistant carcinoid tumors (7–9). Thus, the distinct patterns of phosphorylation and trafficking of sst₅ and sst₂ receptors are likely to regulate differentially the long-term responsiveness of sst₂-expressing and sst₅-expressing tumors. However, the present study only examines phosphorylation and trafficking in cells expressing either sst₂ or sst₅. It would be interesting to know whether sst₂ or sst₅ receptors would be regulated differently in coexpressing cells. Indeed, coexpression of sst₅ receptors has been reported to reduce internalization and desensitization of sst₂ receptors in Chinese hamster ovary K1 cells (28). Thus, it is an attractive hypothesis that the presence of sst₅ receptors would facilitate recycling of sst₂ receptors and thereby regulate the long-term responsiveness to octreotide in carcinoid patients.

In conclusion, we provide direct evidence for agonist-selective phosphorylation of carboxyl-terminal T333. In addition, we identify GRK2-mediated phosphorylation and PP1γ-mediated dephosphorylation of T333 as key regulators of rapid internalization and recycling of the human sst₅ receptor. We also uncover striking differences in the phosphorylation and trafficking patterns of sst₅ and sst₂ receptors, which are likely to be clinically relevant.