Supported by American Heart Foundation SE Affiliate Postdoctoral Fellowship 0425341B. To whom correspondence should be addressed: Dept. of Pharmacology, Emory University School of Medicine, 1510 Clifton Rd., Atlanta, GA 30322. Tel.: 404-727-0363; Fax: 404-727-0365;
‡ Both authors contributed equally to this work. ¶ Supported by American Heart Foundation SE Affiliate Postdoctoral Fellowship 0325259B. * This work was supported in part by National Institutes of Health Grants R01-NS37112 and R01-GM61847 (to J. R. H.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Regulators of G-protein signaling (RGS) proteins act directly on Gα subunits to increase the rate of GTP hydrolysis and to terminate signaling. However, the mechanisms involved in determining their specificities of action in cells remain unclear. Recent evidence has raised the possibility that RGS proteins may interact directly with G-protein-coupled receptors to modulate their activity. By using biochemical, fluorescent imaging, and functional approaches, we found that RGS2 binds directly and selectively to the third intracellular loop of the α1A-adrenergic receptor (AR) in vitro, and is recruited by the unstimulated α1A-AR to the plasma membrane in cells to inhibit receptor and Gq/11 signaling. This interaction was specific, because RGS2 did not interact with the highly homologous α1B- or α1D-ARs, and the closely related RGS16 did not interact with any α1-ARs. The N terminus of RGS2 was required for association with α1A-ARs and inhibition of signaling, and amino acids Lys219, Ser220, and Arg238 within the α1A-AR i3 loop were found to be essential for this interaction. These findings demonstrate that certain RGS proteins can directly interact with preferred G-protein-coupled receptors to modulate their signaling with a high degree of specificity.
Signaling through G-protein-coupled receptors (GPCRs)
The abbreviations used are: GPCR, G-protein-coupled receptor; RGS, regulators of G-protein signaling; NE, norepinephrine; GST, glutathione S-transferase; AR, adrenergic receptor; HA, hemagglutinin; GFP, green fluorescent protein; PBS, phosphate-buffered saline; InsP, inositol phosphate.
1The abbreviations used are: GPCR, G-protein-coupled receptor; RGS, regulators of G-protein signaling; NE, norepinephrine; GST, glutathione S-transferase; AR, adrenergic receptor; HA, hemagglutinin; GFP, green fluorescent protein; PBS, phosphate-buffered saline; InsP, inositol phosphate.
must be tightly regulated in cells to maintain functional specificity. Recent studies have identified a large family of proteins, the Regulators of G-protein Signaling (RGS), which bind Gα-GTP to increase the rate of GTP hydrolysis and rapidly terminate responses (
), it was initially postulated that the specificity of RGS function would be controlled by formation of individual RGS/Gα pairs. However, subsequent studies showed that many RGS proteins, in particular most members of the B/R4 subfamily (RGS1-5, -8, -13, -16, and -18), could nonselectively bind and inhibit Gαi/o and Gαq/11 function in reconstituted systems (
), indicating that RGS proteins and GPCRs are physically and functionally coupled in plants, and suggests that they may have evolved as separate genes in higher organisms allowing for the formation of specific paired complexes. Consistent with this idea, we showed recently (
) that purified RGS2 binds directly to the third intracellular (i3) loops of the Gαq-coupled M1, M3, and M5 muscarinic acetylcholine receptors but not the Gαi-coupled M2 and M4. Binding of RGS2 to the M1 receptor was shown to recruit recombinant RGS2 and to regulate the activity of endogenous Gαq/11 in cell membranes (
). Taken together, these data support the idea that RGS proteins form direct functional complexes with preferred GPCRs in order to modulate the signaling properties of these receptors and their linked G-proteins (
). However, previous work has not yet demonstrated direct RGS interactions with full-length receptors in intact systems, the specific amino acids within a receptor that bind the RGS protein, and the general applicability of this phenomenon across GPCRs.
α1-Adrenergic receptors (ARs) mediate responses to norepinephrine (NE) and epinephrine (
). There are three α1-AR subtypes (α1A, α1B, and α1D) which, like M1 receptors, also activate Gαq and release intracellular Ca2+. α1-ARs play an essential role in the regulation of vascular tone and blood pressure and are an important target for treatment of hypertension (
). Most interestingly, a recent report showed that RGS2 knock-out mice have a strongly hypertensive phenotype, with increased mean arterial pressure, renovascular abnormalities, and persistent constriction of the peripheral vasculature (
). Thus, we examined the possibility that RGS2 might interact specifically with α1-AR subtypes to modulate their signaling. Unexpectedly, we found that RGS2 binds specifically to the i3 loop of the α1A-AR but not the closely related α1B- or α1D-AR. This interaction requires specific amino acids in the α1A-AR i3 loop and results in inhibition of agonist-stimulated responses in intact cells. The findings of this study demonstrate that specific RGS proteins interact directly with preferred GPCRs and that this interaction is essential for controlling RGS specificity and function.
Constructs-GST-α1-i3 Constructs—α1A-i3, α1B-i3, and α1D-i3 loop constructs were originally cloned into the pET41b vector to encode a fusion protein with an N-terminal GST tag and a C-terminal His tag. Each receptor i3 loop was amplified from corresponding regions of the human full-length receptor as follows: α1A, amino acids 213-260; α1B, amino acids 233-280; α1D, amino acids 283-334. α1A and α1D were amplified as BamHI/XhoI fragments, and α1B was amplified as an EcoRI/XhoI fragment. In order to perform pull-down assays with purified His-tagged RGS proteins, fragments encoding the i3 loops of α1A- and α1B-ARs were subcloned further into the pGEX4T vector to eliminate the His tag. α1A was amplified as a BamHI/XhoI fragment, and α1B was amplified as an EcoRI/XhoI fragment and cloned in-frame with an N-terminal GST tag.
α1A/B i3 Loop Chimeras—α1A/B i3 loop chimeras were created by using a nested primer PCR strategy (
) and inserted into the pGEX4T-1 vector. The α1A/B chimera encodes amino acids 214-239 of α1A fused to amino acids 259-280 of α1B. The α1B/A chimera encodes amino acids 234-258 of α1B fused to amino acids 239-260 of α1A. The junction of these chimeras occurs between conserved lysine-asparagine residues.
The α1A/B 1-8 constructs encode single, double, or triple amino acid substitutions of the α1A-i3 sequence with α1B placed into the GST-α1A/B chimeric template. Substitutions were made as denoted below, where the amino acid number corresponds to position of the residue in the context of the full-length α1A-AR receptor sequence: α1A/B-1, E214T/S215T; α1A/B-2, R216K/G217N; α1A/B-3, K219E/S220A; α1A/B-4, L222V/K223M; α1A/B-5, T224K/D225E/K226M; α1A/B-6, D228N; α1A/B-7, E230K/Q231E/V232L; α1A/B-8, R238S. The mutations demonstrating the largest loss in RGS2 binding capacity were then inserted into the full-length α1A-AR in the pDT vector using the Quikchange site-directed mutagenesis kit (Stratagene) for use in functional assays examining RGS2 inhibition of [3H]InsP formation.
RGS Constructs—RGS2-HA, RGS16-HA, N2/RGS16-HA, RGS2-His, RGS16-His, RGS2-GFP, and RGS16-GFP were created as described previously (
α1-AR Constructs—Full-length α1A-AR i3 loop mutant constructs were created using the Quikchange site-directed mutagenesis kit (Stratagene), using HA-α1A-AR in pDoubleTrouble (pDT) as a template. N-terminal HA epitope-tagged human α1-ARs were produced as described previously (
Induction and Purification of GST Fusion Proteins—GST-α1-AR i3 loop fusion proteins (GST-α1A-i3, GST-α1B-i3, GST-α1D-i3, GST-α1A/B-i3, and GST-α1B/A-i3 chimeras and GST α1A/B-i3 mutants) were transformed into BL21(DE3) Escherichia coli and purified as described previously (
Cell Cultures and Transfections—Human embryonic kidney (HEK)293 cells were propagated in Dulbecco's modified Eagle's medium with sodium pyruvate supplemented with 10% heat-inactivated fetal bovine serum, 100 μg/ml streptomycin, and 100 units/ml penicillin at 37 °C in a humidified atmosphere with 5% CO2. Confluent plates were subcultured at a ratio of 1:5 for transfection. HEK293 cells were transfected with 3 μg of DNA of each construct for 24 h using Lipofectamine 2000 transfection reagent (Invitrogen), and cells were used for experimentation 48-72 h after transfection. Wild-type Chinese hamster ovary-K1 cells were grown and maintained in Dulbecco's modified Eagle's medium (CellGro) containing 10% fetal bovine serum (Atlanta Biologicals), 1× nonessential amino acids (CellGro), and penicillin/streptomycin.
Preparation of Cell Lysates—To produce cell lysates containing RGS-HA proteins, wild-type Chinese hamster ovary-K1 cells were grown to 80-90% confluency in 10-cm dishes and transfected by using Lipofectamine 2000 transfection reagent, and cell lysates were harvested as described previously (
). The single modification of this assay involved pull-downs using RGS-His proteins, in which 30 mm imidazole and 80 mm NaCl were included in the buffer to control for differences in the buffers among the purified RGS proteins.
Immunoblots—Immunoblots were performed as described previously (
). In brief, nitrocellulose membranes were incubated in blocking buffer (Tris-buffered saline with 5% milk, 0.5% Tween 20, 0.02% sodium azide) 1 h at room temperature or overnight at 4 °C. Membranes were probed with either mouse anti-HA antibody (Covance) or mouse anti-His (Qiagen) antibodies diluted 1:1000 in blocking buffer for 1-2 h at room temperature. Membranes were washed three times with Tris-buffered saline + 0.1% Tween 20 and then probed with horseradish peroxidase-conjugated goat anti-mouse (Rockland) diluted 1:20,000 in Tris-buffered saline + 0.1% Tween 20. Protein bands were visualized using chemiluminescence and exposed to film.
Laser Confocal Microscopy—Cells transiently transfected with HA- or GFP-tagged constructs were grown on sterile coverslips, fixed for 30 min with 2% paraformaldehyde in 0.1 m phosphate buffer, pH 7.4, and rinsed three times with phosphate-buffered saline (PBS) containing 0.5% normal horse serum (PBS+). For anti-HA immunostaining, fixed coverslips were blocked for 1 h in blocking buffer (PBS containing 1% bovine serum albumin, 5% normal horse serum) containing 0.01% Triton X-100 to permeabilize cells. Anti-HA antibody (1:1000 dilution) was added to coverslips overnight at 4 °C in blocking buffer, washed three times with PBS+, and incubated with rhodamine red-conjugated anti-rabbit IgG secondary antibody (1:500 dilution) for 1 h at room temperature in blocking buffer. Coverslips were washed three times with PBS+ and mounted onto slides using Vectashield mounting medium. Cells were scanned with a Zeiss LSM 510 laser-scanning confocal microscope (Heidelberg, Germany) as described previously (
). For detecting GFP, fluorescein isothiocyanate fluorescence was excited using an argon laser at a wavelength of 488 nm, and the absorbed wavelength was detected for 510-520 nm. For detecting rhodamine fluorescence, a helium-neon laser at a wavelength of 522 nm was used.
Measurement of [3H]InsP Formation—Accumulation of [3H]inositol phosphates (InsPs) was determined in confluent 96-well plates. Transiently transfected HEK293 cells were prelabeled with myo-[3H]inositol for 24 h, and production of [3H]InsP was determined by modification of a protocol described previously (
). After prelabeling, medium containing myo-[3H]inositol was removed, and 1 ml of Krebs-Ringer bicarbonate buffer (in mm: NaCl 120, KCl 5.5, CaCl2 2.5, NaH2PO4 1.2, MgCl2 1.2, NaHCO3 20, glucose 11, Na2EDTA 0.029) containing 10 mm LiCl was gently added to each well. Cells were incubated with or without 100 μm NE for 60 min. The reaction was stopped by addition of 500 μl of methanol, and samples were sonicated for 10 s and centrifuged for 5 min at 10,000 × g. Samples were subjected to anion exchange chromatography to isolate [3H]InsPs, which were quantified by scintillation counting. Percent hydrolysis of total myo-[3H]inositol incorporated into lipid converted into [3H]InsPs was counted and expressed as mean ± S.E. Mean values were compared using the one-sample t test, with a p value less than 0.01 considered significant.
Radioligand Binding—Confluent 150-mm plates were washed with PBS and harvested by scraping. Cells were collected by centrifugation, homogenized with a Polytron, centrifuged at 30,000 × g for 20 min, and resuspended in PBS. Radioligand-binding sites were measured by saturation analysis of specific binding of the α1-AR antagonist radioligand 125I-BE2254 (20-800 pm). Nonspecific binding was defined as binding in the presence of 10 μm phentolamine. The pharmacological specificity of radioligand-binding sites was determined by displacement of 125I-BE2254 (50-70 pm) by NE. Data were analyzed using nonlinear regression (
RGS2 Selectively Associates with the i3 Loop of the α1A-AR—To determine whether selected B/R4 RGS proteins directly associate with α1-ARs, we examined the capacity of closely related RGS2 and RGS16 to associate with the i3 loops of all three subtypes (α1A, α1B, and α1D) by using pull-down assays. As shown in Fig. 1, RGS2 was capable of binding to the α1A-i3, but not to the α1B-i3 or α1D-i3, whereas RGS16 did not associate with any of the three α1-AR subtype i3 loops. To identify a specific RGS2 binding domain within the α1A-i3, we created α1A/B-i3 chimeras encoding amino acids 214-239 of α1A fused to amino acids 259-280 of α1B and α1B/A chimeras encoding amino acids 234-258 of α1B fused to amino acids 239-260 of α1A (Fig. 1B). Chimeras were then fused to GST and tested for their capacity to associate with RGS2-His by using pull-down assays. Most interestingly, we found that RGS2 robustly associated with the GST-α1A/B chimera (Fig. 1C), indicating that the N-terminal half of the α1A-i3 contains an RGS2-binding motif. In contrast, we found RGS2 interacted very weakly or not at all with the GST-α1B/A chimera, suggesting the first 26 amino acids of the i3 loop is predominantly responsible for RGS2 interaction. Therefore, these data indicate that an RGS2-binding motif may be contained within the proximal domain of the α1A-i3.
RGS2 Co-localizes at the Plasma Membrane with α1A-ARs in HEK293 Cells—To determine whether RGS2 associates with α1A-ARs in a cellular context, we co-transfected GFP-tagged RGS proteins with HA-tagged α1-ARs into HEK293 cells, and we examined their cellular localization by using confocal microscopy. However, to ensure that these effects are not a result of receptor and/or RGS overexpression, we performed radioligand binding experiments using 125I-BE2254 on cells transiently expressing HA-α1A-AR and HA-α1B-ARs, and we found that both constructs express between 100 and 300 fmol/mg protein (data not shown), which is well within the physiological range for expression of these receptors in native systems (
). In addition, we found that when titrating the concentration of GFP-tagged RGS constructs to be used in transfection from 0.5 to 6 μg of cDNA per 30-mm plate, 3 μg of construct resulted in ∼60% transfection efficiency with the majority of the cells exhibiting low to moderate fluorescence (data not shown). In agreement with previous studies, RGS2 (
). However, upon co-transfection with HA-α1A-ARs, RGS2-GFP was primarily localized at the plasma membrane (Fig. 2B, upper panels), whereas RGS2-GFP remained sequestered within the nucleus when co-transfected with HA-α1B-ARs (Fig. 2B, middle panels). In addition, RGS16-GFP cellular localization was unchanged upon co-expression with HA-α1A- (Fig. 2B, lower panels) or HA-α1B-ARs (data not shown), indicating this association occurs in an RGS-specific manner. Therefore, these experiments support our findings from the pull-down assays demonstrating that RGS2 can selectively associate with α1A-ARs.
) indicates that RGS proteins are recruited to the plasma membrane when co-expressed with GPCR in an agonist-independent manner. To examine the effects of receptor ligand binding on α1A-AR/RGS2 co-localization, HEK293 cells transiently co-expressing HA-α1A-ARs and RGS2-GFP were stimulated with norepinephrine, niguldipine, and prazosin, respectively. Addition of 100 nm of the α1A-AR subtype-selective antagonist niguldipine for 30 min did not alter the co-localization of α1A-AR/RGS2 at the plasma membrane (Fig. 2C, upper panel). Incubating cells with the putative α1-AR inverse agonist prazosin (100 nm, 30 min) also did not alter the plasma membrane localization of α1A-AR/RGS, although the proteins appeared to be clustered and nonoverlapping in their distribution (Fig. 2C, middle panel). However, stimulation of cells with 10 μm NE for 30 min resulted in marked α1A-AR internalization with an apparent disruption of the α1A-AR-RGS2 complex (Fig. 2C, lower panel). This was also observed following 5 and 15 min of stimulation with 10 μm NE and with 5, 15, and 30 min of stimulation with 1 μm NE (data not shown). Therefore, these data suggest that agonist stimulation results in α1A-AR internalization, which is followed by subsequent RGS2 dissociation from the receptor and redistribution to the cytosol.
RGS2 Selectively Inhibits α1A-AR Functional Responses in HEK293 Cells—We next examined if RGS2 can selectively regulate α1A-AR functional responses, by transiently transfecting α1-ARs into HEK293 alone and in combination with RGS2 proteins and assaying for NE-stimulated [3H]InsP formation. Both HA- and GFP-tagged RGS2 significantly inhibited [3H]InsP formation by α1A-ARs in response to 100 μm NE by 60.6 ± 3.3 and 61.4 ± 5.3%, respectively (Fig. 3, left). Transfection with pEGFP or pcDNA3.1 vector alone had no effect on α1A-AR signaling. However, HA- and GFP-tagged RGS2 had no significant effect on α1B-AR-stimulated [3H]InsP formation (Fig. 3, right). Combined with our previous findings, these data indicate that RGS2 selectively associates with α1A-ARs at the plasma membrane to facilitate uncoupling of the receptor with G-protein-mediated functional responses.
Effect of RGS2/α1A-AR Association on Agonist Binding Affinity—It is generally accepted that agonists display multiple affinity states for GPCRs, as a result of G-protein binding to GPCR i3 loops. Typically, binding of the G-protein induces low affinity agonist binding interactions with the GPCR, and uncoupling of the G-protein by GTP decreases agonist affinity by >10-fold. In our studies, we find that RGS2 can directly associate with the α1A-i3. We therefore tested whether RGS2 affected agonist affinity for binding to the α1A-AR. To examine this, we harvested membranes from HEK293 cells stably expressing wild-type α1A-ARs, which were then preincubated for 30 min at 4 °C with concentrations of purified RGS2-HA reported previously to be sufficient for maximal association with M1 receptors in isolated Chinese hamster ovary cell membranes (
). After 30 min, the affinity of the nonselective adrenergic receptor agonist NE was determined using 125I-BE2254 competition radioligand binding. As reported in Table I, NE bound with low affinity to α1A-ARs when expressed alone. However, preincubation with 10, 30, or 100 nm RGS2 caused no significant change in affinity, suggesting that RGS2 binding to the α1A-AR does not affect ligand-receptor interactions.
Table IEffect of RGS2/α1A-AR association on agonist affinity
The N Terminus of RGS2 Determines Selectivity of Receptor Association—Previously, we and others have shown the N-terminal domains of RGS proteins are required for promoting association with the Gα subunits and with GPCRs (
). However, the functional significance of this interaction was not examined in intact cells. Therefore, we examined the capacity of a chimera containing the N-terminal portion of RGS2 fused to the RGS domain and C-terminal portion of RGS16 (N2/RGS16-HA) to associate with and inhibit the signaling of α1A-ARs (Fig. 4A). In pull-down assays, we found RGS2-HA and N2/RGS16-HA selectively associated with α1A-i3 GST-fusion proteins, whereas RGS16-HA did not (Fig. 4A). In HEK293 cells transiently co-transfected with HA-α1A-ARs alone or with RGS constructs, co-expression with RGS2-HA resulted in significant inhibition of α1A-AR stimulation of [3H]InsP formation, whereas co-expression with RGS16-HA caused no significant decrease in the α1A-AR maximal response (Fig. 4B). However, co-expression with N2/RGS16-HA caused a decrease in NE-stimulated [3H]InsP formation to the level observed with RGS2 (Fig. 4B). Taken together, these data suggest that the N-terminal region of RGS2 is responsible for association with and inhibition of α1A-AR function, through a direct association with the i3 loop.
Identification of Amino Acids within the α1A-i3 Responsible for Binding of RGS2—Thus far, we have determined that RGS2 associates with the α1A-AR through a direct association with the proximal half of the i3 loop. Next, we initiated studies to identify specific amino acids within the α1A-i3 that are responsible for promoting this interaction. In Fig. 1, we showed RGS2 binds the α1A/B but not the α1B/A-i3 chimera. By comparing the sequence homology between the α1A-i3 and α1B-i3 (Fig. 5A), nonhomologous amino acids in the proximal half were identified between the receptors to target for substitution mutation, and used to create a series of α1A-i3 mutants in which specific amino acids in the α1A-i3 were replaced with the corresponding amino acids in the α1B-i3, using the α1A/B as a template. A total of eight mutant α1A-i3 constructs were created using PCR, each involving between 1 and 3 amino acid substitutions (Fig. 5A). Constructs were then expressed as GST fusion proteins and were used for pull-down assays with purified RGS2-His. Of the eight α1A/B-i3 mutants examined, the constructs containing a Lys219-Ser220 to Glu-Ala (A/B3), Leu222-Lys223 to Val-Met (A/B4), and single Arg238 to Ser (A/B8) mutation demonstrated the most severe loss in RGS2 binding (Fig. 5B).
To determine whether the loss of binding correlated with a decrease in RGS2 inhibition of α1A-AR functional responses, the Lys-Ser/Glu-Ala, Leu-Lys/Val-Met, and Arg/Ser mutations were introduced into full-length α1A-ARs via site-directed mutagenesis, using the full-length α1A-AR in pDT vector as a template. Full-length α1A-ARs carrying the Lys-Ser/Glu-Ala, Leu-Lys/Val-Met, and Arg/Ser mutations were then transiently transfected into HEK293 alone or in combination with RGS2-HA and assayed for NE-stimulated [3H]InsP formation. In comparison to full-length α1A-ARs (Bmax = 321 ± 32 fmol/mg protein), each of the three α1A-AR mutants demonstrated relatively equal (Bmax values, Lys-Ser/Glu-Ala = 401 ± 15, Arg/Ser = 300 ± 24 fmol/mg protein) or higher (Bmax Leu-Lys/Val-Met = 710 ± 45 fmol/mg protein) binding site densities. As shown in Fig. 6, RGS2 caused a significant decrease in the efficacy of NE for stimulating α1A-AR-mediated phosphatidylinositol hydrolysis (Fig. 6A). However, α1A-ARs containing either the Lys-Ser/Glu-Ala (Fig. 6B) or Arg/Ser mutations (Fig. 6C) were rendered insensitive to RGS2, indicating that these mutations abrogated the direct association between RGS2 and α1A-ARs. However, α1A-ARs containing the Leu-Lys/Val-Met mutation remained susceptible to RGS2 inhibition of functional responses (Fig. 6D) indicating that in the context of the full-length receptor in a cellular environment, these amino acids alone are not required for RGS2 association with receptor. Taken together, these data suggest that RGS2 associates with the α1A-AR through a direct interaction within the proximal half of the α1A-i3 and that three amino acids within this domain (Lys219, Ser220, and Arg238) appear to be critical for this interaction.
Many RGS proteins, particularly those of the B/R4 family, show little selectivity in inhibiting G-protein signaling when assayed as purified proteins in reconstituted systems, but they appear to have a high degree of specificity in intact cells (
) suggest this may be due to the capacity of GPCRs to recruit RGS proteins to the plasma membrane and thus regulate the specificity of RGS function, although these studies provide only indirect evidence in support of this hypothesis. Our recent work (
) demonstrates that RGS2 binds directly and selectively to the intracellular third loop of the M1 muscarinic receptor to modulate receptor and Gq/11 signaling in cell membranes. However, this and other previous work has not demonstrated direct RGS interactions with full-length receptors in intact systems, the specific amino acids within a receptor that binds RGS proteins, or the general applicability of this phenomenon across GPCRs. In this study, we demonstrate that RGS2 directly associates with α1A-ARs through an interaction between the N-terminal domain of RGS2 and three specific amino acids in the proximal half of the α1A-AR i3 loop. Through this association, α1A-ARs recruit RGS2 to the plasma membrane, resulting in inhibition of agonist-stimulated responses. In contrast, RGS2 was incapable of directly associating with or regulating the function of the closely related α1B-AR. Thus, we demonstrate here that this direct RGS interaction is selective for specific Gq/11-linked receptors and that this interaction is necessary to confer RGS specificity of function.
Previously, RGS2 has been shown to selectively block Gαq/11 function in reconstituted systems (
). Most unexpectedly, RGS2 interacted specifically with the α1A-but not the closely related α1B- or α1D-AR subtypes. This specificity appeared to be due to RGS2 associating with specific amino acids in the α1A-AR i3 loops, as replacement with the corresponding amino acids in the α1B-AR completely abrogated the capacity of RGS2 to bind and inhibit α1A-AR agonist-stimulated responses. These data demonstrate that binding motifs located within GPCR i3 loops are responsible for RGS binding and functional specificity. In addition, these data demonstrate that an RGS protein will not necessarily interact with all GPCRs linked to a common signaling pathway.
Our findings show that RGS2 is selectively recruited to the plasma membrane by the unstimulated α1A-AR to regulate receptor and G-protein signaling. These findings are consistent with our previous findings with M1 muscarinic receptors (
) showing specific membrane recruitment of certain RGS proteins by unstimulated receptors. Taken together, these findings suggest that receptors and RGS proteins can form preferred functional pairs and predict a model where GPCR and RGS are prebound prior to signal initiation. By forming a complex with a specific GPCR at the plasma membrane, the RGS is positioned to modulate the linked G-protein upon agonist activation. In this way, the receptor determines RGS protein/G-protein coupling and functional specificity. Consistent with this model, we found that RGS16 is capable of blocking receptor-mediated Gq/11 signaling only when it contained the N terminus of RGS2 and is capable of binding α1A-AR. Further studies will be needed to confirm these models.
Based on a number of recent studies, it is now apparent that RGS proteins play a prominent role in regulating GPCR function in the cardiovascular system (
). Therefore, the hypertensive phenotype observed in RGS2 knock-out mice may be due to an increased activity of the α1A-AR in the peripheral vasculature. Thus, based on the results of this and previous studies, RGS proteins appear to be attractive new therapeutic targets for development of novel pharmaceutical agents (
This study presents evidence that RGS2 binds directly to α1A-ARs to modulate their function. This interaction occurs with a high degree of specificity through identified domains within the receptor and the RGS protein. Thus, we propose that RGS proteins form stable, functional complexes with preferred GPCRs to selectively modulate the signaling functions of those receptors and linked G-proteins. Such direct interactions are likely to play significant roles in the still poorly understood specificity of RGS actions in vivo.
We thank Dr. Howard Rees and Dr. Allan Levey for help with confocal microscopy studies.