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Research Article

Amendment history:

Dopamine 5 receptor mediates Ang II type 1 receptor degradation via a ubiquitin-proteasome pathway in mice and human cells

Hewang Li1, Ines Armando1, Peiying Yu1, Crisanto Escano1, Susette C. Mueller2, Laureano Asico1, Annabelle Pascua1, Quansheng Lu1, Xiaoyan Wang1, Van Anthony M. Villar1, John E. Jones1, Zheng Wang1, Ammasi Periasamy3, Yuen-Sum Lau4, Patricio Soares-da-Silva5, Karen Creswell6, Gaétan Guillemette7, David R. Sibley8, Gilbert Eisner9, Robin A. Felder10 and Pedro A. Jose1


1Department of Pediatrics and
2Department of Oncology, Lombardi Comprehensive Cancer Center, Georgetown University Medical Center, Washington, DC, USA.
3Keck Center for Cellular Imaging and Biology Department, University of Virginia, Charlottesville, Virginia, USA.
4Department of Pharmacological and Pharmaceutical Sciences, College of Pharmacy, University of Houston, Houston, Texas, USA.
5Institute of Pharmacology and Therapeutics, Faculty of Medicine of Porto, Porto, Portugal.
6Flow Cytometry and Cell Sorting Center, Lombardi Comprehensive Cancer Center, Georgetown University Medical Center, Washington, DC, USA.
7Department of Pharmacology, Faculty of Medicine and Health Sciences, Université de Sherbrooke, Sherbrooke, Quebec, Canada.
8Molecular Neuropharmacology Section, National Institute of Neurological Disorders and Stroke (NINDS), NIH, Bethesda, Maryland, USA.
9Department of Medicine, Georgetown University Medical Center, Washington, DC, USA.
10Department of Pathology, University of Virginia Health Sciences Center, Charlottesville, Virginia, USA.

Address correspondence to: Pedro A. Jose, Department of Pediatrics, Georgetown University Medical Center, Washington, DC 20007, USA. Phone: (202) 444-8675; Fax: (202) 444-7161; E-mail: pjose01@georgetown.edu.

First published May 8, 2008
Submitted: August 16, 2007; Accepted: March 19, 2008.

Hypertension is a multigenic disorder in which abnormal counterregulation between dopamine and Ang II plays a role. Recent studies suggest that this counterregulation results, at least in part, from regulation of the expression of both the antihypertensive dopamine 5 receptor (D5R) and the prohypertensive Ang II type 1 receptor (AT1R). In this report, we investigated the in vivo and in vitro interaction between these GPCRs. Disruption of the gene encoding D5R in mice increased both blood pressure and AT1R protein expression, and the increase in blood pressure was reversed by AT1R blockade. Activation of D5R increased the degradation of glycosylated AT1R in proteasomes in HEK cells and human renal proximal tubule cells heterologously and endogenously expressing human AT1R and D5R. Confocal microscopy, Förster/fluorescence resonance energy transfer microscopy, and fluorescence lifetime imaging microscopy revealed that activation of D5R initiated ubiquitination of the glycosylated AT1R at the plasma membrane. The regulated degradation of AT1R via a ubiquitin/proteasome pathway by activation of D5R provides what we believe to be a novel mechanism whereby blood pressure can be regulated by the interaction of 2 counterregulatory GPCRs. Our results therefore suggest that treatments for hypertension might be optimized by designing compounds that can target the AT1R and the D5R.

Introduction

Hypertension is associated with sodium retention caused by decreased renal sodium excretion, which is tightly regulated by both natriuretic and antinatriuretic hormones. Dopamine and Ang II are among those hormones that have counterregulatory effects on cellular signal transduction, production of reactive oxygen species, renal sodium excretion, and blood pressure via renal dopamine D1-like receptors (dopamine 1 receptor [D1R] and D5R subtypes), D2-like receptors (D2R, D3R, and D4R subtypes), and Ang II type 1 receptors (AT1Rs) (14). The D5R is widely expressed in the rodent kidney, specifically in the proximal and distal tubules, cortical collecting ducts, medullary ascending limbs of Henle, and arterioles, but not in the glomeruli, juxtaglomerular cells, or macula densa (5). The AT1R is also widely expressed in the kidney, specifically in the proximal and distal tubules, cortical collecting ducts, medullary ascending limbs of Henle, arterioles, glomeruli, juxtaglomerular cells, and macula densa (2, 68).

Recent evidence shows that fenoldopam (Fen), a D1-like receptor agonist, decreases AT1R protein expression in rat renal proximal tubules (RPTs) and in vascular smooth muscle cells (9). Gene knockdown studies using antisense oligonucleotides that target either the D1R or D5R transcript point to D5R, not D1R, as the negative regulator of AT1R protein expression in human RPT cells (10). Furthermore, renal AT1R protein expression is greater in D5R knockout (Drd5–/–) mice than in wild-type (Drd5+/+) littermates (9), and renal D5R protein expression is increased in Agtr1a–/– mice, underscoring the important downregulatory function of D5R on AT1R protein expression, and vice versa. Moreover, Drd5–/– mice are hypertensive and salt sensitive (11), while Agtr1a–/– mice have decreased blood pressure (12), indicating the importance of dopamine receptors (5) and regulators (e.g., GPCR kinase 4; ref. 13) in the regulation of blood pressure.

In this report, we investigated the in vivo and in vitro interaction between D5R and AT1R and explored the mechanisms involved in this interaction. We showed that the high blood pressure of Drd5–/– mice, which was associated with increased AT1R protein expression, was normalized by AT1R blockade. Activation of the D5R decreased AT1R protein level by increasing AT1R degradation in proteasomes via a ubiquitin/proteasome pathway; the degrading action was initiated by the ubiquitination of the glycosylated AT1R at the plasma membrane. Stimulation of the D5R dissociated the interaction of the D5R with the AT1R and increased the proteasomal degradation of the ubiquitinated AT1Rs. We believe our results demonstrate a novel paradigm of GPCR-mediated cellular responses and regulation of degradation of one GPCR by another and moreover provide a novel mechanism for the regulation of blood pressure by the dopaminergic system.

Results

Elevated blood pressure in Drd5–/– mice is normalized by an AT1R antagonist. To test whether the increased expression of the AT1R in the absence of the D5R is functionally significant, Drd5+/+ and Drd5–/– mice were treated via intraperitoneal injection with vehicle or the AT1R antagonist losartan daily for 5–7 days. Consistent with previous observations (9, 11), both blood pressure (Figure 1A) and AT1R protein expression (Figure 1B) were significantly increased in Drd5–/– mice relative to Drd5+/+ littermates (n = 7; P < 0.01). Radioligand binding autoradiography showed that AT1R expression increased in the renal cortical tubules but not in the glomeruli of Drd5–/– mice (Supplemental Table 1; supplemental material available online with this article; doi: 10.1172/JCI33637DS1). Losartan did not change AT1R protein expression in either Drd5+/+ or Drd5–/– mice (Figure 1B), but it decreased blood pressure (systolic, diastolic, and mean arterial pressures) in Drd5–/– but not Drd5+/+ mice (Figure 1A), which indicates that the lack of the D5R increases the effect of the AT1R on blood pressure and that increased expression of the AT1R, among other factors, is important in the pathogenesis and maintenance of hypertension in Drd5–/– mice.

Effect of the AT1R antagonist losartan on blood pressure and AT1R and reninFigure 1

Effect of the AT1R antagonist losartan on blood pressure and AT1R and renin protein expression in Drd5+/+ and Drd5–/– mice. Vehicle (Veh) or losartan (Los) was administered via intraperitoneal injection every day for 5–7 days in 6-month-old mice. (A) Blood pressure was measured directly via the femoral artery under pentobarbital anesthesia. Losartan decreased blood pressure in Drd5–/– but not Drd5+/+ mice. SBP, systolic blood pressure; DBP, diastolic blood pressure; MAP, mean arterial pressure. n = 7. *P < 0.05 versus all other groups, factorial ANOVA, Holm-Sidak test. (B and C) Whole homogenates (30 μg) of kidneys isolated from A were electrophoresed (SDS-PAGE gel) and immunoblotted with anti-AT1R (B) or anti-renin (C) antibody. Amounts of glycosylated AT1R and renin proteins relative to actin were quantified by densitometry. The lanes for the renin blot were run on the same gel but were noncontiguous. n = 3–5. *P < 0.05 versus vehicle-treated Drd5+/+, factorial ANOVA, Holm-Sidak test. Data are mean ± SEM.

We next investigated whether renin, another component of the renin-angiotensin system (RAS), was altered in Drd5–/– mice. As shown in Figure 1C, renal renin protein expression was not significantly altered in Drd5–/– mice compared with their Drd5+/+ littermates. As expected, losartan increased renal renin protein expression in Drd5+/+ mice (n = 3–5; P < 0.01); this effect was not observed in Drd5–/– mice (n = 3–5; P = 0.307).

Elevated blood pressure in Drd5–/– mice is not associated with alterations in renal or plasma catecholamines. We previously reported that high blood pressure of anesthetized Drd5–/– mice was, at least in part, due to increased central nervous sympathetic tone, based on the greater percentage reduction in blood pressure with α2-adrenergic blockade but similar percentage reduction in blood pressure after adrenalectomy in Drd5–/– mice relative to their wild-type littermates (11). Here, renal and plasma levels of norepinephrine, epinephrine, and l-DOPA were not different between the 2 mouse strains (Supplemental Table 2). However, the renal dopamine level increased in Drd5–/– mice relative to their wild-type littermates (Supplemental Table 2). Therefore, the elevated blood pressure in Drd5–/– mice was not caused by increased peripheral catecholamines or a deficiency in renal dopamine levels.

Activation of D5R decreases AT1R protein expression in vitro. Renal cross-transplantation experiments have previously shown that the kidney contributes about 50% of blood pressure regulation (14). The RPT and thick ascending limb of Henle are the sites of increased sodium reabsorption in human essential polygenic hypertension (15), whereas the distal nephron is the site of increased sodium transport in monogenic hypertension (16). Therefore, we next studied the effect of D1-like receptor stimulation on AT1R protein expression in human RPT cells, which endogenously expressed D1R (data not shown), D5R (Supplemental Figure 2), and AT1R (Supplemental Figure 2). In human RPT cells, Fen (1 μM for 20 min) decreased AT1R protein expression (Figure 2). The effect of Fen on AT1R protein expression was observed for the approximately 70–90 kDa band, but not for the approximately 40–60 kDa band; presumably, the former represents the glycosylated form of AT1R (17, 18). The observed change in AT1R expression was specifically caused by D1-like receptor stimulation, because cotreatment with SCH23390, a D1-like receptor antagonist that had no effect by itself, abrogated the decrease in AT1R protein expression (Figure 2). These data indicate that a D1-like receptor, presumably the D5R (9, 10), negatively regulates AT1R protein expression in human RPT cells.

Effect of Fen on AT1R protein expression.
   Figure 2

Effect of Fen on AT1R protein expression. Immortalized human RPT cells were incubated with vehicle (lane 1), Fen (1 μM for 20 min; lane 2), or SCH 23390 (1 μM for 50 min) in the absence (lane 3) or presence (lane 4) of Fen. AT1R protein expression in total cell lysates was determined by immunoblotting with anti-AT1R (top panel): 2 bands were observed, denoted a and b, which presumably represent the glycosylated and nonglycosylated forms of the AT1R, respectively (17, 18). Immunoblotting for actin (bottom panel) showed equal sample loading in each lane. Shown is 1 immunoblot from 3 experiments. The effect of vehicle, Fen, or SCH 23390 in the presence or absence of Fen on AT1R protein expression was also quantified. n = 3. *P < 0.05 versus vehicle, ANOVA, Student-Newman-Keuls test. Data are mean ± SEM.

There are no pharmacological reagents that can distinguish the 2 subtypes of D1-like receptors, D1R and D5R (1, 19). Therefore, we transfected D5R into HEK293 cells, which express no endogenous dopamine receptors (20, 21). To investigate the mechanism of the regulation of AT1R protein expression by the D5R, we transfected human full-length AT1R tagged at the C terminus with EGFP into HEK293 cells expressing the D5R. Flow cytometry (Supplemental Figure 1A) and immunoblot analyses (Supplemental Figure 1B) of the AT1R-EGFP transfectants confirmed the stable heterologous expression of AT1R-EGFP in D5R HEK293 cells. Radioligand binding assays (Supplemental Figure 1C) using 3H-SCH 23390 and 125I-Sar1 Ang II as ligands for the D1-like receptors (here, only the D5R) and the AT1R, respectively, also showed the expression of both the D5R and the AT1R. Ang II increased the phosphorylation of ERK1/2 in a dose-dependent manner (Supplemental Figure 1D), indicating that the EGFP tag did not impair AT1R function in the transfected cells.

We next tested whether activation of the D5R decreased AT1R protein expression in the transfected cells (designated AT1R/D5R HEK293 cells). Fen decreased the approximately 100–120 kDa band, which presumably represents the glycosylated AT1R plus the EGFP tag, but not the approximately 65–85 kDa AT1R band, which presumably represents the nonglycosylated AT1R plus the EGFP tag, as early as 20 minutes after treatment (Figure 3A). Because the D1R is not expressed in HEK293 cells (20, 21), these results indicate that the effect of Fen on AT1R protein expression is solely the result of D5R activation.

Effect of D5R stimulation on the expression of the glycosylated form of theFigure 3

Effect of D5R stimulation on the expression of the glycosylated form of the AT1R protein. (A) AT1R/D5R HEK293 cells were incubated with Fen (1 μM) for the indicated times; total cell lysates were electrophoretically separated, transferred onto nitrocellulose membranes, and immunoblotted with anti-GFP (band a, ~100–120 kDa; band b, ~65–85 kDa) to detect AT1R protein expression. Immunoblotting for actin was used to monitor the sample protein loading. n = 3. *P < 0.05 versus 0 min, ANOVA, Student-Newman-Keuls test. (B) Protein samples (50 μg) from AT1R/D5R HEK293 cells were incubated with 10 mU PNGase F at 37°C for the indicated times, electrophoretically separated, transferred onto nitrocellulose membranes, and immunoblotted with anti-GFP antibody. Band S is sensitive to PNGase F and corresponds to the glycosylated form of the AT1R (band a). Band R is resistant to PNGase F and corresponds to the nonglycosylated form of the AT1R (band b). The ratio of band S intensity to band R intensity was determined by densitometry in 3 separate experiments. n = 3. *P < 0.05 versus 0 h, ANOVA, Student-Newman-Keuls test. Data are mean ± SEM.

Treatment with PNGase F (Figure 3B) greatly decreased the intensity of the approximately 100–120 kDa AT1R band, which indicates that this band represents the glycosylated form of the receptor. Thus, the D5R only affects the expression of the glycosylated species of AT1R. Moreover, the rapid D5R-mediated decrease of this band, but not the approximately 65–85 kDa AT1R band (Figure 3A), indicates that the inhibitory effect does not result from the downregulation of transcription or translation, but rather may be the result of increased AT1R protein degradation.

Characterization of D5R-mediated AT1R degradation. To characterize the D5R-mediated AT1R turnover, we performed pulse-chase experiments by [35S] methionine/cysteine (Met/Cys) labeling of AT1R/D5R HEK293 cells treated with vehicle, Fen, or Ang II. Stimulation of the D5R with Fen resulted in degradation of the AT1R more rapid than that observed with vehicle treatment, but slower than that with Ang II treatment (Figure 4). Unlike Ang II, which increased degradation of both the glycosylated and the nonglycosylated AT1R, Fen increased the degradation of only the glycosylated AT1R (Figure 4A), and the amount of the approximately 100–120 kDa AT1R band decreased as a function of time (0.365 + 0.618t/22.2), which suggests a first-order decay.

Characterization of AT1R degradation by [35S] metabolic labeling.
   Figure 4

Characterization of AT1R degradation by [35S] metabolic labeling. AT1R/D5R HEK293 cells were treated with vehicle, Ang II (100 nM for 30 min), or Fen (1 μM for 2 h), pulsed with [35S] Met/Cys, and then chased with cold amino acids for the indicated times. The cell lysates were immunoprecipitated with anti-GFP antibody, and the immunocomplexes were subjected to SDS-PAGE. Dried gels were exposed to X-ray films. (A) Autoradiographs of AT1R degradation with vehicle, Ang II, and Fen treatment. (B) Quantification of AT1R degradation. Values are mean ± SEM of arbitrary density units normalized to the time-0 value (n = 3). The decrease in AT1R protein in the presence of vehicle followed a first-order exponential curve, with a half-life of 127.6 min. The decrease in AT1R protein with Ang II or Fen treatment also followed a first-order exponential curve; treatment with Ang II decreased the half-life of AT1R protein to 16.8 min, while treatment with Fen resulted in a half-life of 37.7 min. P < 0.05, Fen versus vehicle and versus Ang II, repeated-measures ANOVA. No error bar is visible when the SEM is smaller than the symbol.

Protein degradation in mammalian cells occurs primarily in either lysosomes or proteasomes (22, 23). Spontaneous or Ang II–induced AT1R degradation occurs in lysosomes (24, 25). To explore the sites of D5R-mediated AT1R degradation in AT1R/D5R HEK293 cells, we examined the colocalization of the AT1R with marker proteins for proteasomes or lysosomes using laser scanning confocal microscopy. Under basal conditions, a substantial fraction of AT1Rs (43.47%) were localized at the plasma membrane (Supplemental Figure 3 and Supplemental Table 3), consistent with the previous observation that the mouse AT1AR mainly localizes at the plasma membrane (26). Lysosome-associated membrane protein 1 (LAMP 1), a marker of lysosomes, and p44S10, a marker of proteasomes, were scattered throughout the cytoplasm with much less expression at the plasma membrane (Supplemental Figure 3). In the cytoplasm, there was some colocalization of the AT1R and LAMP 1 (23.19%) and of the AT1R and p44S10, albeit to a lesser degree (11.26%; Supplemental Figure 3 and Supplemental Table 4). Stimulation with Fen resulted in considerable internalization of AT1Rs with patches and clusters of AT1R fluorescence in both plasma membrane and cytoplasm. Fen treatment increased the colocalization of AT1R with p44S10 from 11.26% to 56.12%, whereas the treatment did not affect the percentage of colocalization between AT1Rs and LAMP 1 (Supplemental Figure 3 and Supplemental Table 4). In contrast, treatment with Ang II increased the colocalization between AT1Rs and LAMP 1 without affecting that between AT1R and p44S10 (Supplemental Figure 3). These results strongly suggest that upon activation, the D5R directs the degradation of the AT1R to proteasomes and not to lysosomes. The Fen-induced changes in the intracellular distribution of the AT1R did not occur in the cells incubated with SCH23390 prior to Fen treatment (Supplemental Figure 3 and Supplemental Tables 3 and 4), which indicates that the effect of Fen was through the activation of the D5R.

D5R-mediated AT1R degradation occurs in proteasomes, but not lysosomes. To investigate further the D5R-mediated AT1R degradation in AT1R/D5R HEK293 cells, we used clasto-lactocystin β-lactone (CLBL), a proteasome inhibitor, and chloroquine (Chlor), a lysosome inhibitor, in [35S] Met/Cys metabolic experiments. Fen-stimulated AT1R degradation was greatly reduced in the presence of CLBL but not Chlor; the half-life of the AT1R in the presence of CLBL was 149.5 min (decay rate, 0.537), while it was 43.8 min (decay rate, 0.665) in the presence of Chlor (Figure 5A). Morphological observation showed that CLBL, but not Chlor, abrogated the Fen-stimulated colocalization of AT1R and p44S10 (Figure 5B and Supplemental Table 4). In the presence of CLBL, image analysis showed that 42.40% of AT1R resided in the plasma membrane in spite of the presence of Fen (Supplemental Table 3). Flow cytometry data also showed that CLBL, but not Chlor, inhibited the Fen-stimulated degradation of the AT1R (Figure 5C). Chlor alone slightly increased the AT1R fluorescence (Figure 5C), consistent with previous observations that spontaneous degradation of the AT1R occurs in lysosomes (24, 25). Again, inhibition of proteasome but not lysosome activity abrogated D5R-mediated AT1R degradation, which confirmed that this degradation occurs in proteasomes.

Fen-stimulated AT1R degradation occurs in proteasomes but not lysosomes.
  Figure 5

Fen-stimulated AT1R degradation occurs in proteasomes but not lysosomes. (A) AT1R/D5R HEK293 cells treated with Fen (1 μM for 2 h) and either Chlor (100 μM for 3 h) or CLBL (4 μM for 3 h) were pulsed with [35S] Met/Cys and chased with cold amino acids for the indicated times. Shown is 1 autoradiograph of each treatment. Data (mean ± SEM) followed a first-order exponential curve for both treatments. The AT1R protein half-life was 43.8 min for Fen plus Chlor and 149.5 min for Fen plus CLBL. n = 3. (B) Sub­cellular distribution of AT1R in AT1R/D5R HEK293 cells treated as in A. Green, AT1R-EGFP; red, lysosomes or proteasomes (detected by Alexa Fluor 633–conjugated anti–LAMP 1 or anti-p44S10); yellow, colocalization. Scale bars: 10 μm; 2.5 μm (insets). (C) AT1R/D5R HEK293 cells were treated with vehicle, Fen (1 μM for 6 h), Chlor (100 μM for 10 h), CLBL (4 μM for 10 h), Chlor plus Fen, or CLBL plus Fen. Nontransfected D5R HEK293 cells are shown as a control. Right: MFI of AT1R-EGFP. n = 3. *P < 0.05, **P < 0.001 versus vehicle, ANOVA, Student-Newman-Keuls test. Data are mean ± SEM. (D) Human RPT cells were treated with vehicle, Ang II, or Fen; cell lysates were incubated with control or proteasome-affinity beads for 4 hours at 4°C. After washing, the matrix was suspended in SDS sample buffer, separated by SDS-PAGE, and transferred onto nitrocellulose. Membranes were immunoblotted for either AT1R or p44S10. Experiments were repeated twice with similar results.

To corroborate the biochemical and morphological studies indicating that exogenously and endogenously expressed AT1Rs are directed to proteasomes after Fen stimulation, proteasomes were isolated from human RPT cells treated with vehicle, Ang II, or Fen. Using proteasome affinity beads, AT1Rs were identified in proteasomes isolated from Fen-treated human RPT cells, but not in those isolated from vehicle- and Ang II–treated cells (Figure 5D), which indicates that Fen stimulation directs endogenously expressed AT1R to proteasomes in human RPT cells.

AT1R interacts with ubiquitin and is ubiquitinated at the plasma membrane. Ubiquitin (Ub) modification plays an important role in targeting proteins for degradation (23, 2732). To determine the role of ubiquitination in the degradation of the AT1R, we first studied the colocalization of Ub (and ubiquitinated proteins), p44S10, and the AT1R in vehicle- and Fen-treated cells (Figure 6A). In vehicle-treated cells, Ub and p44S10 were scattered throughout the cytoplasm, and minimal — if any — colocalization was observed among the 3 proteins (Figure 6A). In contrast, Fen treatment induced significant colocalization of the AT1R and p44S10, the AT1R and Ub, Ub and p44S10, and the AT1R, Ub, and p44S10 at the plasma membrane (Figure 6A and Supplemental Table 5).

The AT1R is ubiquitinated, and the ubiquitination of the AT1R is initiated Figure 6

The AT1R is ubiquitinated, and the ubiquitination of the AT1R is initiated at the plasma membrane. AT1R/D5R HEK293 cells were treated with vehicle or Fen (1 μM for 5 min). (A) Immunostaining for p44S10 or Ub (clone P4D1). Distance calibration was based on objective and zoom of the images taken: 1 pixel equals 0.133 μm. Green, EGFP-tagged AT1R; red, p44S10; blue, Ub; yellow, colocalization of AT1R and p44S10 (green and red); cyan, colocalization of AT1R and Ub (green and blue); magenta, colocalization of p44S10 and Ub (red and blue); white, colocalization of AT1R, Ub, and p44S10 (red, green, and blue). Scale bars shown for vehicle-treated cells also apply to Fen-treated cells. (B) The image of AT1R-EGFP (as donor fluorophore) was used for drawing ROIs, which were divided into plasma membrane (rectangles) and cytoplasm (ovals) to determine the spatial interaction between the AT1R and Ub (clone P4D1, Alexa Fluor 555; as acceptor fluorophore). The pure FRET and 2-dimensional distribution images of energy transfer efficiency (E%) were generated as described in Methods. Graphs show mean ± SEM energy transfer efficiency and distance (r) of ROIs in the plasma membrane (Mem) and cytoplasm (Cyt) in the corresponding AT1R images processed from 6–8 cells. *P < 0.05 versus vehicle, Student’s t test. Distances beyond experimental limitations (>90 υ) are given a 0 value, indicating no occurrence of FRET.

To validate the morphological findings observed with confocal microscopy, Förster/fluorescence resonance energy transfer (FRET) microcopy (33) was used to detect the interaction between the AT1R and endogenous Ub in AT1R/D5R HEK293 cells. Fen stimulation markedly increased the energy transfer of the AT1R and Ub at the plasma membrane from 3.6% ± 1.2% in unstimulated cells to 34.1% ± 2.4% (Figure 6B). At the same time, the energy transfer had no obvious increase in the cytoplasm, which indicates that Fen stimulation increased the AT1R and Ub interaction initially at the plasma membrane but not in the cytoplasm.

Fluorescence lifetime imaging microscopy (FLIM) allows quantitative monitoring of protein interaction by measuring the change in lifetime of the donor molecule, which does not require any spectral bleed-through correction and is independent of change in fluorophore concentration and excitation intensity (34). To verify the above observations obtained with FRET microscopy, FLIM was used to measure the interaction between the AT1R and Ub at the plasma membrane by monitoring the quench time (τ) of the AT1R (as a donor) in the presence or absence of Ub (as an acceptor). As shown in Figure 7A, the average τm of the AT1R was 2.32 ns in the absence of the acceptor and 2.33 ns in vehicle-treated cells in the presence of the acceptor. Fen stimulation shortened the average τ1 of the AT1R to 1.68 ns because of the occurrence of FRET between the AT1R and Ub, which suggests their interaction upon Fen stimulation. There was no interaction between them in the basal state, consistent with the confocal and FRET microscopy observations.

Glycosylation is necessary for the ubiquitination of AT1R at the plasma memFigure 7

Glycosylation is necessary for the ubiquitination of AT1R at the plasma membrane. (A) AT1R/D5R HEK 293 cells were treated with vehicle or Fen (1 μM for 5 min). Shown are FLIM images, histograms, and decay graphs of cells treated in the absence (top) or presence (middle and bottom) of acceptor (Ub). The left shift (arrow) of AT1R lifetime in Fen-treated samples demonstrated the occurrence of FRET. The lifetime of AT1R (donor) in the absence of acceptor was 2.32 ± 0.2 ns; it was 2.33 ± 0.2 ns in the presence of acceptor with vehicle treatment, indicating no energy transfer. With Fen treatment in the presence of acceptor, the donor lifetime had 2 peaks: the first was quenched at 1.68 ± 0.2 ns (τ1) because of the occurrence of FRET, with a corresponding energy transfer efficiency of 27.6% ± 3.7%; the second was 2.24 ± 0.3 ns (τ2), which represented the unquenched AT1R. The FLIM patterns observed in the basal state and after Fen were similar in 4 different experiments. Quantification was performed in 16–30 cells from 1 of the 4 experiments. Data are mean ± SEM. (B) Human RPT cells were treated with vehicle, Fen (1 μM for 5 min), tunicamycin (Tunica; 10 μg/ml for 12 h), or tunicamycin plus Fen. Cell membrane fractions isolated by biotinylation were immunoprecipitated with normal mouse Ig M/G (lane 1), normal rabbit IgG (lane 2), anti-AT1R mAb (lane 3), or anti-D5R antibody (lane 4) and immunoblotted with anti-Ub (clone FK1) or anti-AT1R mAb. (Ub)n, polyubiquitin chain; HC, heavy chain.

To determine directly whether the AT1R is ubiquitinated, human RPT (Figure 7B) and AT1R/D5R HEK293 (Supplemental Figure 4) plasma membrane fractions were isolated by biotinylation, immunoprecipitated with anti-AT1R or anti-GFP, and immunoblotted with Ub antibody (clone FK1). Fen caused the polyubiquitination of both endogenous (Figure 7B) and heterologously overexpressed (Supplemental Figure 4) AT1R, which was not observed in the basal state. The Fen-stimulated ubiquitination of AT1R was similar in the presence (RPT cells) or absence (AT1R/D5R HEK293 cells) of the D1R, which indicates that the D1R was not involved in D5R-mediated AT1R ubiquitination. The Fen-stimulated polyubiquitination of the endogenous AT1R was prevented by tunicamycin treatment (Figure 7B), indicating that glycosylation of the AT1R was necessary for its ubiquitination at the plasma membrane. Coimmunoprecipitation of the endogenous AT1R with the D5R in vehicle-treated cells, but not in Fen-treated cells, suggested that the AT1R interacted with the D5R in the basal, unstimulated state and that Fen dissociated their interaction (Figure 7B). Failure to detect coimmunoprecipitation of the AT1R with the D5R after tunicamycin treatment indicated that the interaction of the AT1R and the D5R in the basal state required glycosylation of the AT1R and/or the D5R.

Discussion

Our results demonstrate what we believe to be a novel mechanism of the regulation of the AT1R protein expression, which is physiologically important in vivo. The D5R mediates the degradation of glycosylated AT1R through the Ub/proteasome pathway, a process initiated at the plasma membrane, in contrast to either spontaneous or Ang II–mediated degradation of the AT1R, which occurs in lysosomes.

Ubiquitination plays important roles in the regulation of proteins destined for lysosome or proteasome degradation (28, 35). A number of GPCRs in mammalian cells have been reported to be ubiquitinated, such as the β2-adrenegic receptor (36), chemokine receptor CXCR4 (37, 38), platelet-activating factor receptor (39), and protease-activated receptors 1 and 2 (40, 41). However, to our knowledge, no previous studies have suggested that the AT1R is ubiquitinated following its stimulation. Indeed, the agonist-mediated endocytosis of the rodent AT1AR does not require ubiquitination of the receptor (42).

Lysine is the target amino acid for ubiquitination. The human AT1R contains many lysine residues, especially in its intracellular domains: in 19% (10 of 53) of its carboxy terminus, 20% (4 of 20) of its third intracellular loop, 7% (1 of 15) of its second intracellular loop, and 20% (2 of 10) of its first intracellular loop. Even though ubiquitination is not required for rodent AT1AR internalization either in the basal state or following Ang II stimulation in CHO cells (42), human AT1R ubiquitination can still occur under certain conditions. Our results showed that following short-term D5R stimulation, Ub accumulated at sites where the AT1R was located at the plasma membrane (Figure 6, A and B, and Figure 7A), where it was initially ubiquitinated (Figure 7B). The ubiquitinated AT1Rs were then directed to proteasomes for subsequent degradation (Figures 57).

N-glycosylation is a common posttranslational modification of GPCRs and plays roles in the formation of their active conformation, targeting to the cell surface and intracellular signaling (43). Human bradykinin B2 receptor requires N-glycosylation for its maturation at the cell surface (44); certain sites of β2-adrenergic receptor N-glycosylation can alter receptor targeting to the degradation pathway (45). Both dopamine receptors (46, 47) and the AT1R (17, 18) are N-glycosylated. Glycosylation is required for the D5R to be functionally expressed at the cell surface (46). N-glycosylation of the AT1R is important for its proper folding, targeting, and cell surface display (18). Our results showed that glycosylation was required for the interaction of AT1R with D5R (Figure 7B), which is consistent with previous observation that glycosylation is important for their homologous (44) or heterologous (47) interaction with their binding partners to accomplish a series of intracellular functions. Glycosylation may also serve as a signal for AT1R ubiquitination upon Fen stimulation and subsequent degradation in proteasomes (refs. 4850, Figure 7B, and Supplemental Figure 4), which is consistent with a previous observation that N-linked high-mannose oligosaccharides recognized by Ub ligase lead to the ubiquitination of N-glycosylated proteins (49). Therefore, we speculate that ubiquitination of the glycosylated AT1R expressed at the plasma membrane initiates its degradation in proteasomes (Figures 6 and 7). Additionally, glycosylation may protect the nascent AT1R from proteolytic degradation (6, 7, 48). However, the protection is superseded by activation of the D5R, resulting in targeting of the receptor to proteasomes for degradation (Figures 6 and 7).

Both the D5R and the AT1R are GPCRs that are important in the regulation of sodium balance and blood pressure. The RAS regulates blood pressure by renal and nonrenal mechanisms (2, 6, 7, 14, 51). The paracrine regulation of sodium transport in the proximal tubule by the RAS is exerted via several angiotensin receptor subtypes (AT1R, AT2R, and AT4R) (2, 6, 7). The activation of AT1Rs by Ang II increases sodium transport, whereas the activation of AT2R and AT4R decreases sodium reabsorption. Under physiologic conditions, the major effect of Ang II is to stimulate sodium transport via the AT1R (2, 6, 7). In the hypertensive state, the AT1R function is increased, which may or may not be associated with increased expression (2, 68). However, the mechanism underlying this phenomenon in RPTs is not completely clear. Assessment of the in vivo expression levels of D5R and AT1R following dopaminergic or AT1R stimulation may clarify this issue. Dopamine and its receptors have been reported to participate in the regulation of the RAS, including Ang II receptors (5, 9, 52, 53). D1-like receptors have been reported to increase the AT2R (53) and decrease the AT1R (9, 52) expression in RPTs. The D1-like receptor subtype regulating the expression of a particular angiotensin receptor subtype has not been established.

Second-generation Drd5–/– mice in a mixed B129Sv and C57BL/6 background (11, 21) and F6 Drd5–/– mice in a C57BL/6 (>98% congenic) background (9, 21) have high blood pressure (9, 11, 21) and increased AT1R protein expression (9). Renal or plasma catecholamine levels (Supplemental Table 2) could not explain the increased blood pressure and AT1R protein expression observed in Drd5–/– mice. Renal renin protein expression was not different between Drd5–/– and Drd5+/+ littermates (Figure 1C). Renin is mainly synthesized and secreted by juxtaglomerular cells, which express AT1R and D1R, but not D5R (11). Because D5R is not expressed in juxtaglomerular cells, one would expect no change in AT1R expression or alteration in renin levels via the AT1R/renin feedback loop. Although D1R can increase renin secretion, D1R protein expression in the brain and kidney is not altered by knocking out the gene encoding D5R (11). This indicates that the juxtaglomerular AT1R/renin feedback loop is not altered by knocking out the D5R gene. Consistent with previous reports (54), AT1R blockade with losartan treatment increased renal renin expression in Drd5+/+ mice. Interestingly, this effect was not observed in Drd5–/– mice, the mechanism of which remains to be determined. Nonetheless, our results showed that the increased blood pressure in Drd5–/– mice was probably not caused by increased production of renin and/or Ang II.

The results of the present study show that the hypertension resulting from the absence of the gene encoding D5R is caused, at least in part, by the increase in AT1R protein expression, because of the absence of the negative counterregulatory effect of the D5R on the AT1R. Activation of the D5R initiated AT1R ubiquitination, which directed the AT1R to proteasomal degradation (Figures 57). Glycosylation was required for the ubiquitination of the AT1R (Figure 7). These results provide evidence for a mechanism by 2 posttranslational modifications crucial in the regulation of the AT1R, a class B GPCR, mediated by the D5R, a class A GPCR. The negative regulation of the AT1R expression by the D5R has physiological consequences, because chronic administration of the AT1R blocker losartan normalized the elevated blood pressure of Drd5–/– mice (Figure 1A). Inhibition of the RAS is an important component of the treatment of essential hypertension, but the control of hypertension remains suboptimal (55). Our results showing that the antihypertensive effect of D1-like receptors counteracted the prohypertensive effects of the RAS, in part by D5R-mediated degradation of the AT1R, provides opportunities for designing specific drugs that can target the AT1R and/or the D5R for the optimization of hypertension therapy.

Methods

Antibodies and reagents; cell lines and transfection; radioligand binding; radioligand binding autoradiography; biotinylation and immunoprecipitation; deglycosylation; proteasome isolation; flow cytometry; and determination of catecholamine levels. See Supplemental Methods.

Drd5–/– mice and blood pressure measurement. The animal protocols were reviewed and approved by the Georgetown University Animal Care and Use Committee. For details, see Supplemental Methods.

Pulse-chase. AT1R/D5R HEK293 cells were labeled by 35S-labeled Met/Cys as described previously (56). Briefly, cells were starved in Met/Cys-free DMEM (Invitrogen) containing 10% dialyzed FBS for 2 h and pulsed for 3 h with medium containing 300 μCi/ml 35S-labeled Met/Cys. Cycloheximide (10 μg/ml) was added to inhibit de novo protein synthesis and then chased for varying durations in full culture medium containing 3 mM unlabeled Met/Cys. At the end of each individual chase period, the cells were washed with PBS 3 times to remove unbound label. Cell pellets were lysed with lysis buffer, and the AT1R was immunoprecipitated using anti-GFP antibody. Immunoprecipitated protein complexes were eluted with protein sample buffer at 85°C for 15 minutes and resolved by SDS-PAGE. Dried gels were exposed to X-ray films for autoradiography. The quantified AT1R bands were normalized to the time-0 value. The mean and SEM of 3 experiments were plotted, and the exponential decays were calculated (OriginPro; OriginLab Corp.). The decay rate constants and the half-lives of the AT1R protein, which represent the rates of AT1R degradation, were calculated based on individual decay equations.

Confocal immunofluorescence microscopy. AT1R/D5R HEK293 cells were fixed with 4% paraformaldehyde in PBS for 20 min at room temperature. After washing with PBS, the fixed cells on coverslips were incubated overnight at 4°C with Alexa Fluor 633–conjugated (Invitrogen) polyclonal rabbit LAMP 1 (2 μg/ml) or polyclonal goat p44S10 antibody (5 μg/ml), or Alexa Fluor 546–conjugated (Invitrogen) monoclonal mouse Ub antibody (2 μg/ml). Coverslips were mounted in SlowFade mounting medium (Invitrogen) and sealed onto glass slides. Samples were imaged using an Olympus Fluoview FV300 laser scanning confocal microscope equipped with a ×60/1.4 NA objective. Quantitative analysis was conducted by using MetaMorph 6.1 (Molecular Devices). Background was subtracted from each image, after which the images were thresholded to identify specific protein fluorescence. Whole cells, plasma membrane, or cytoplasmic areas were identified by creating regions of interest (ROIs). Fluorescence in each (integrated intensity) was determined and expressed as percent of total fluorescence for cytoplasm and plasma membrane. For determination of colocalization, the function “Colocalization” in MetaMorph software was used where percent overlap (integrated intensity measurement) was determined for AT1R over LAMP 1, or AT1R over p44S10, Ub over AT1R, and Ub over p44S10.

FRET microscopy and data processing. The fluorophore pairs used for FRET imaging in this study were AT1R-EGFP (as donor dipole) and Alexa Fluor 555 (as acceptor dipole; Invitrogen) conjugated with Ub antibody. Seven images were acquired for each FRET analysis, as described previously (33), with an Olympus Fluoview FV300 laser scanning confocal microscope equipped with a ×60/1.4 NA objective, argon (488 nm) and HeNe (543 nm) laser, emission filters 515/50 nm and 590 nm LP. Either single-labeled donor and acceptor samples or double-labeled samples were acquired under the same condition throughout the image collection. The uncorrected FRET images (uFRET) were acquired by donor excitation in acceptor channel, which contained pure FRET (pFRET) and contaminations from both donor and acceptor spectral bleed-through (SBT). pFRET images were generated using a previously described algorithm (33) for pixel-by-pixel removal of donor and acceptor SBT on the basis of matched fluorescence levels between the double-labeled specimen and single-labeled reference specimens.

ROIs were selected in the uFRET images (33). In this study, we used image e (donor excitation in donor channel of double-labeled specimen) as the reference image for selection of ROIs to separate plasma membrane and cytoplasm fluorescence. The percentage of energy transfer (E%) images was processed on a pixel-by-pixel basis by using the equation E% = 1 – {Ida/(Ida + pFRET × [Pd/Pa] × [Sd/Sa] × [Qd/Qa])}; where Pd and Pa (photo multiplier tube gain of donor and acceptor channels, respectively) set to the same when images were acquired; Sd and Sa (spectral sensitivity of donor and acceptor channels, respectively) were provided by the manufacturer; Qd and Qa (donor and acceptor quantum yield, respectively) were measured by spectrofluorometer as described previously (57); Ida is the image of donor excitation in donor channel of double labeling samples after removing of background; and pFRET is the processed or corrected FRET. The calculation of distance of donor and acceptor (r) was based on the following previously described equation (33): r = R0{(1/E)–1}1/6, where E is the efficiency of energy transfer. Förster’s distance (R0) in this study was 67.5 (33).

FLIM. Cell preparation and fluorophores of samples were performed as described above. Time-domain FLIM was performed with a 120-s acquisition in a 2-photon BioRad Radiance 2100 microscope with ×60 NA 1.4 objective lens at the M.W. Keck Center for Cellular Imaging of the University of Virginia, as described previously (34). The AT1R was fused with EGFP as the donor fluorophore, and the acceptor fluorophore was Ub (clone P4D1), conjugated with Alexa Fluor 555. The single exponential decay of lifetime was processed in the donor alone samples, and the double exponential decay of lifetime was performed in the presence of acceptor fluorophores. Donor fluorophores (AT1R-EGFP) were excited at 790 nm by a femtosecond pulse from a coherent ti:sapphire laser system. Emissions filtered with a center of 515 nm were collected by a high-speed photomultiplier tube (MCP R3809; Hamamatsu), and fluorescence lifetime imaging capability was incorporated, with addition of a fast time-correlated single photon counting acquisition board (SPC 730; Becker and Hickl). Fluorophore lifetimes were fitted to 1- or 2-exponential decay curves on SpcImage 2.60 (Becker and Hickl). The non-FRETing population, measured as the donor lifetime in the absence of an acceptor, is represented as τ2; the FRETing population (τ1) is represented in the figures. The energy transfer efficiency is calculated as 1 – (τDAD), where τDA and τD are the donor excited state lifetimes in the presence and absence, respectively, of the acceptor. Cell images and lifetime distributions are representative examples. The FLIM patterns observed in the basal state and after Fen were similar in 4 different experiments. Quantification was performed in 16–30 cells from 1 of the 4 experiments.

Statistics. Results are expressed as mean ± SD or SEM as indicated. Significant differences among groups were determined by factorial or repeated ANOVA (Holm-Sidak or Student-Newman-Keuls test) for multiple comparisons and Student’s t test for 2 comparisons. P < 0.05 was considered statistically significant (SigmaStat 3.0; SPSS Inc.).

Supplemental data

View Supplemental data

Acknowledgments

We are grateful to Tamas Balla (National Institute of Child Health and Human Development, NIH, Bethesda, Maryland, USA) for critically reading the manuscript and to Guillermo Palchik and Michelle Lombard for their assistance with confocal microscopy and flow cytometry, respectively. Our special thanks go to Ye Chen (University of Virginia), who helped with FRET and FLIM microscopy. These studies were supported in part by NIH grants HL23081, DK39308, HL68686, and DK52612 (to P.A. Jose); HL074940 (to R.A. Felder); CA51008 (to S.C. Mueller); and RR15768-01 (to S.C. Mueller and K. Creswell).

Footnotes

Nonstandard abbreviations used: AT1R, Ang II type I receptor; Chlor, chloroquine; CLBL, clasto-lactocystin β-lactone; D5R, dopamine 5 receptor; Fen, fenoldopam; FLIM, fluorescence lifetime imaging microscopy; FRET, Förster/fluorescence resonance energy transfer; LAMP 1, lysosome-associated membrane protein 1; Met/Cys, methionine/cysteine; RAS, renin-angiotensin system; ROI, region of interest; RPT, renal proximal tubule; τ, quench time; Ub, ubiquitin.

Conflict of interest: The authors have declared that no conflict of interest exists.

Citation for this article:J. Clin. Invest.118:2180–2189 (2008). doi:10.1172/JCI33637

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