Molecular determinants of A2AR–D2R allosterism: role of the intracellular loop 3 of the D2R

Authors

  • Víctor Fernández-Dueñas,

    1. Unitat de Farmacologia, Facultat de Medicina, Departament de Patologia i Terapèutica Experimental, Universitat de Barcelona, L'Hospitalet de Llobregat, Barcelona, Spain
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  • Maricel Gómez-Soler,

    1. Unitat de Farmacologia, Facultat de Medicina, Departament de Patologia i Terapèutica Experimental, Universitat de Barcelona, L'Hospitalet de Llobregat, Barcelona, Spain
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  • Kenneth A. Jacobson,

    1. Molecular Recognition Section, Laboratory of Bioorganic Chemistry, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland, USA
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  • Santhosh T. Kumar,

    1. Molecular Recognition Section, Laboratory of Bioorganic Chemistry, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland, USA
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  • Kjell Fuxe,

    1. Department of Neuroscience, Karolinska Institutet, Stockholm, Sweden
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  • Dasiel O. Borroto-Escuela,

    1. Department of Neuroscience, Karolinska Institutet, Stockholm, Sweden
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  • Francisco Ciruela

    Corresponding author
    • Unitat de Farmacologia, Facultat de Medicina, Departament de Patologia i Terapèutica Experimental, Universitat de Barcelona, L'Hospitalet de Llobregat, Barcelona, Spain
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Address correspondence and reprint requests to Francisco Ciruela, Unitat de Farmacologia, Facultat de Medicina, Departament Patologia i Terapèutica Experimental, Universitat de Barcelona, L'Hospitalet de Llobregat, Barcelona 08907, Spain. E-mail: fciruela@ub.edu

Abstract

In the CNS, an antagonistic interaction has been shown between adenosine A2A and dopamine D2 receptors (A2ARs and D2Rs) that may be relevant both in normal and pathological conditions (i.e., Parkinson's disease). Thus, the molecular determinants mediating this receptor–receptor interaction have recently been explored, as the fine tuning of this target (namely the A2AR/D2R oligomer) could possibly improve the treatment of certain CNS diseases. Here, we used a fluorescence resonance energy transfer-based approach to examine the allosteric modulation of the D2R within the A2AR/D2R oligomer and the dependence of this receptor–receptor interaction on two regions rich in positive charges on intracellular loop 3 of the D2R. Interestingly, we observed a negative allosteric effect of the D2R agonist quinpirole on A2AR ligand binding and activation. However, these allosteric effects were abolished upon mutation of specific arginine residues (217–222 and 267–269) on intracellular loop 3 of the D2R, thus demonstrating a major role of these positively charged residues in mediating the observed receptor–receptor interaction. Overall, these results provide structural insights to better understand the functioning of the A2AR/D2R oligomer in living cells.

Abbreviations used
AGT

alkyltransferase

APEC

2-[[2-[4-[2-(2-aminoethyl)-aminocarbonyl]ethyl]phenyl]ethylamino]-5′-N-ethyl-carboxamidoadenosine

FRET

fluorescence resonance energy transfer

HEK

Human embryonic kidney

IL3

third intracellular loop 3

RIPA

radio immunoprecipitation assay

The existence of oligomeric complexes comprising adenosine and dopamine receptors (i.e., A2AR/D2R) has been postulated to be relevant for proper striatal function both in normal and pathological conditions (Fuxe et al. 2010). For instance, these A2AR/D2R oligomers have been implicated in the control of striatal processes such as motor activity control, motor learning, or some forms of associative and visual learning (Bornstein and Daw 2011). Interestingly, the A2AR–D2R interaction occurs in the somatodendritic area of GABAergic enkephalinergic striatal neurons (Cabello et al. 2009; Ferre et al. 2007a,b; Fuxe et al. 1998, 2003). At this level, the coexistence of reciprocal antagonistic interactions between these receptors has been further described, namely an intramembrane interaction in which the A2AR mediates inhibition of the D2R, thus modulating neuronal excitability and neurotransmitter release, and an interaction at the level of adenylate cyclase (AC), in which the D2R inhibits A2AR-mediated protein phosphorylation and gene expression (Shindou et al. 2002; Svenningsson et al. 2000). Thus, the rationale to study the precise mechanism behind these A2AR–D2R molecular and functional interactions is well suited as the A2AR/D2R oligomer has been postulated to be responsible for the neuronal signal integration coming from two different neurotransmitter systems (i.e., dopaminergic and adenosinergic) (Ferre et al. 2007b).

Importantly, in recent years the existence of the A2AR/D2R heteromer has been revealed in living cells, while in native tissue the results are still not conclusive, by biochemical (i.e., co-immunoprecipitation) and biophysical (i.e., resonance energy transfer-based) approaches (Agnati et al. 2005; Cabello et al. 2009; Canals et al., 2003; Genedani et al. 2005; Kamiya et al. 2003). In addition, computerized modeling and techniques of pull down and mass spectrometry have shown the heteromerization between the A2AR and the D2R to depend on a Coulombic interaction between the third intracellular loop (IL3) of the D2R and the C-terminal tail of the A2AR (Canals et al., 2003; Ciruela et al. 2004; Woods and Ferre 2005; Woods et al. 2005). Thus, two positively charged arginine rich motifs (215VLRRRRKRV223 and 265EVRRRNV271) located at the N-terminal part of the D2R-IL3 may interact with two distinct negatively charged motifs from the C-terminal tail of the A2AR. These motifs are the 388HELKGVCPEPPGLDDPLAQDGAVGS412 domain, which contains two adjacent aspartic acid residues, or the 370SAQEpSQGNT378 domain, which contains a phosphorylatable serine residue (S374), and thus their interaction with the D2R forms electrostatic bonds of covalent-like strength (Canals et al., 2003; Ciruela et al. 2004; Woods and Ferre 2005; Woods et al. 2005).

Interestingly, we recently demonstrated that the point mutation of serine 374 to alanine (S374A) reduced A2AR/D2R heteromerization and blocked the allosteric modulation of the D2R by the A2AR (Borroto-Escuela et al. 2010a,b), a result that was further confirmed (Navarro et al. 2010). Also, when the S374A point substitution was accompanied by mutation of the two negatively charged aspartates on the A2AR C-terminal tail (D401A/D402A, see above), a synergistic reduction in the physical A2AR/D2R interaction and the loss of antagonistic allosteric modulation over the A2AR/D2R interface were observed (Borroto-Escuela et al. 2010a,b). Similarly, it was recently shown the importance of the serine residue on the present A2AR–D2R interaction (Azdad et al. 2009). Thus, by means of competitive peptides mimicking the serine-containing epitope it was possible to preclude, performing perforated-patch-clamp recordings on brain slices, the ability of the A2AR to counteract the effects of D2R activation (Azdad et al. 2009). In addition, by using synthetic transmembrane (TM) α-helical peptides of the D2R, the role of helical interactions within the A2AR/D2R heteromeric TM interface was also recently explored. Thus, TMs-IV and V of the D2R may play a critical role in the A2AR/D2R heteromeric interface as incubation with peptides corresponding to these domains significantly reduced the ability of the A2AR and D2R to heteromerize. Also, the incubation with TM-IV or TM-V peptides blocked the allosteric modulation normally found in the A2AR/D2R heteromer (Borroto-Escuela et al. 2010a,b). Overall, the above-mentioned results further corroborated the existence of an electrostatic interaction between the C-terminal tail of the A2AR and IL3 of the D2R. On the other hand, while the A2AR-mediated allosteric effects on the D2R have been object of numerous studies, the reciprocal interaction is a less studied aspect of this antagonistic receptor–receptor interaction. Hence, in the present work, we aimed to study the D2R-mediated allosteric modulation of the A2AR functionality within the framework of A2AR/D2R oligomers, and to demonstrate that this receptor–receptor allosterism is mostly mediated by two regions rich in positive charges located within IL3 of the D2R.

Materials and methods

Plasmid constructs

We created an A2AR sensor (A2ARCFP) to perform fluorescence resonance energy transfer (FRET) experiments with the fluorescent agonist MRS5424, a nucleoside labeled strategically with Alexa Fluor 532. To this end, a pcDNA3.1 vector (Invitrogen, Carlsbad, CA, USA) containing (from 5′ to 3′) a signal peptide that targets the receptor to the cell membrane, the fluorescent protein Cyan Fluorescence Protein (CFP), and the parathyroid hormone receptor type 1 (PTHR1) was used as a template (kindly provided by Dr. J. P. Vilardaga, University of Pittsburgh, Pittsburgh, PA, USA). Thus, the cDNA encoding the A2AR (Cabello et al. 2009) was amplified by a polymerase chain reaction using the primers FA2ABam (5′-CGCGGATCCATGCCCATCATGGGCTCC-3′) and RA2AXho (5′-CGCCTCGAGTCAGGACACCCTGTCTCC-3′), and subcloned into the BamHI/XhoI sites replacing the PTHR1 in the above-mentioned template vector.

The mutations at the IL3 of the D2R were introduced into the cDNA encoding the wild-type D2R cloned in pEYFP-N1 (i.e., D2RYFP) (Clontech, Heidelberg, Germany) (Cabello et al. 2009). Thus, two regions rich in positive charges (amino acids 217–222 and 267–269) within the IL3 were sequentially mutated to alanine by using the QuickChange site-directed mutagenesis kit (Stratagene Europe, Amsterdam, The Netherlands) and following the manufacturer's instructions. Accordingly, the 217–222 region was first mutated from RRRRKR to AAAAKA, and thereafter, the obtained 217–222 mutant D2R cDNA was used as a template for the second set of mutations (267R–269R–267A–269A). The mutagenic primers were designed using Stratagene's web-based program: F217–222A (5′-ATCAAGATCTACATTGTCCTCGCCGCAGCCGCCAAGGCAGTCAACACCAAACGCAGCAGC-3′) and R217–222A (5′-GCTGCTGCGTTTGGTGTTGACTGCCTTGGCGGCTGCGGCGAGGACAATGTAGATCTT-3′) for the first set of mutations, and F267–269A (5′-TGGGAGTTTCCCAGTGAAGCGGGCGGCAGTGGTGCAGGAGGCTGC-3′) and R267–269A (5′-GGCAGCCTCCTGCACCACTGCCGCCGCGTTCACTGGGAAACTCCC-3′) for the second. The resulting mutated D2R construct, D2RmutYFP, was verified by DNA sequencing with BIGDYE® terminator V3.1 cycle sequencing kit (Applied Biosystems Hispania S.A., Alcobendas, Spain).

Finally, to visualize the presence of the D2R during our real-time FRET experiments, we tagged D2R constructs at their N-terminus with the O6-alkylguanine-DNA alkyltransferase (AGT), which is a 24-kDa protein that acts as a suicide enzyme to covalently transfer modifications from DNA bases onto it. Interestingly, once the AGT is fused to the protein of interest, it is possible to stain it by means of fluorescent AGT substrates, which if non-permeable allow the identification of cell surface proteins over periods of hours because of the fact that the complex formed is extremely stable (Ciruela et al. 2010). To this end, a pcDNA3.1 vector containing (from 5′ to 3′) the signal peptide of the metabotropic glutamate 5 receptor, the AGT protein and adenosine A1 receptor was used as a template (kindly provided by Dr. J. P. Pin, Université de Montpellier and Institut de Génomique Fonctionnelle, Montpellier, France). Thus, the cDNA encoding the D2R and the D2Rmut were amplified by polymerase chain reaction using the primers FD2SNAP (5′-GCCGCTCGAGGATCCACTGAATCTGTCCTGG-3′) and RD2SNAP (5′-GCCGAAGCTTTCAGCAGTGGAGGATCTTCAGGAAGGCC-3′), and subcloned into the XhoI/HindIII sites replacing the A1 receptor in the above-mentioned template vector.

Synthesis of the A2AR fluorescent agonist

APECAlexa532 (MRS5424) is a functional agonist at the adenosine A2AR, in which the fluorescent dye Alexa Fluor 532 is covalently attached to a functionalized A2AR agonist 2-[[2-[4-[2-(2-aminoethyl)-aminocarbonyl]ethyl]phenyl]ethylamino]-5′-N-ethyl-carboxamidoadenosine (APEC). Briefly, similar to the method previously described for a related APEC conjugate (Brand et al. 2008), MRS5424 was synthesized as follows: Alexa Fluor 532 carboxylic acid, N-succinimidyl ester (1.0 mg, 1.38 μmol) was dissolved in anhydrous DMF (200 μL). Freshly prepared sodium tetraborate labeling buffer (0.1 M, 1 mL, pH 8.5) and APEC (1.12 mg, 2.07 μmol), which was dissolved in anhydrous DMF (200 μL), were added. The reaction mixture was protected from light and after stirring for 18 h, the mixture was diluted with H2O (600 μL) and purification was performed by HPLC with a Luna 5 μ RP-C18(2) semipreparative column (250 × 10.0 mm; Phenomenex, Torrance, CA, USA) under the following conditions: flow rate of 2 mL/min; 10 mM triethylammonium acetate (TEAA)-CH3CN from 100 : 0 (v/v) to 70 : 30 (v/v) in 30 min. The homogeneous product was isolated in the triethylammonium salt form with an HPLC retention time of 13.5 min. Analytical purity of this conjugate was checked using a Hewlett-Packard 1100 HPLC equipped with a Zorbax SB-Aq 5 μm analytical column (50 × 4.6 mm; Agilent Technologies Inc, Palo Alto, CA, USA). Mobile phase: linear gradient solvent system: 5 mM TBAP (tetrabutylammonium dihydrogenphosphate)-CH3CN from 80 : 20 to 40 : 60 in 13 min; the flow rate was 0.5 mL/min (retention time 9.08 min). Peaks were detected by UV absorption with a diode array detector at 254, 275, and 280 nm, and the yield of MRS5424 was 0.67 mg (31%). ESI-HRMS m/z 1150.4142 [M + H]+, C55H63N11O13S2·H+: Calcd. 1150.4127).

Cell culture, transfection, and membrane preparation

Human embryonic kidney (HEK293) cells were grown in Dulbecco's modified Eagle's medium (Sigma-Aldrich) supplemented with 1 mM sodium pyruvate, 2 mM l-glutamine, 100 U⁄mL streptomycin, 100 mg⁄mL penicillin and 5% (v⁄v) fetal bovine serum at 37°C and in an atmosphere of 5% CO2. HEK293 cells growing in 25-cm2 flasks or six-well plates containing 18 mm coverslips were used for western blot analysis and fluorescence imaging, respectively, were transiently transfected with the cDNA encoding the specified proteins using TransFectin Lipid Reagent (Bio-Rad Laboratories, Hercules, CA, USA). Membrane suspensions from transfected HEK293 cells were obtained as described previously (Burgueno et al. 2003, 2004).

cAMP determinations

The functionality of both the D2RYFP and D2RmutYFP and their allosteric modulation on activity of the A2AR were assessed by means of a dual luciferase reporter assay (Promega, Stockholm, Sweden). In brief, upon co-transfection of a plasmid containing the CRE fused to firefly luciferase, it is possible to indirectly detect variations of cAMP, as the expression of the reporter leads to increased levels of luminescence, which are directly proportional to newly generated cAMP. Thus, cells were co-transfected with plasmids corresponding to the following constructs as follows: 1 μg firefly luciferase-encoding experimental plasmid (pGL4-CRE-luc2p; Promega), 1 μg of D2RYFP or D2RmutYFP expression vectors, 1 μg of A2ARCFP expression vector (only in the allosteric studies), and 50 ng Renilla luciferase-encoding internal control plasmid (phRG-B; Promega). Approximately 36 h post-transfection, after the cells were treated for 4 h with forskolin (Sigma-Aldrich, St. Louis, MO, USA), quinpirole (Sigma-Aldrich), CGS21680 (Tocris Bioscience, Ellisville, MI, USA), and/or ZM 241385 (Tocris Bioscience), they were harvested with passive lysis buffer (Promega), and the luciferase activity of cell extracts was determined in a FLUOStar Optima plate-reader (BMG Labtech, Durham, NC, USA) using a 30-nm bandwidth excitation filter at 535 nm. Firefly luciferase was measured as firefly luciferase luminescence over a 15 s reaction period. The luciferase values were normalized against Renilla luciferase luminescence values.

Immunoprecipitation, gel electrophoresis, and immunoblotting

For immunoprecipitation, membrane suspensions of transiently transfected HEK293 cells were solubilized in ice-cold radio immunoprecipitation assay (RIPA) buffer (50 mM Tris-HCl, pH 7.4, 100 mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.2% sodium dodecyl sulfate, and 1 mM EDTA) for 30 min on ice. The solubilized preparation was then centrifuged for 30 min at 13200 rpm at 4°C. The supernatant (1 mg/mL) was processed for immunoprecipitation, each step of which was conducted with constant rotation at 0–4°C. The supernatant was incubated overnight with 1 μg of mouse anti-A2AR monoclonal antibody (clone 7F6-G5-A2; Millipore, Temecula, CA, USA). Then, a suspension of protein A cross-linked to agarose beads (40 μL, Sigma-Aldrich) was added, and the mixture was incubated overnight. The beads were washed twice with ice-cold RIPA buffer, twice with ice-cold RIPA buffer diluted 1 : 10 in (Tris-buffered saline; 50 mM Tris-HCl pH 7.4, 100 mM NaCl), and once with Tris-buffered saline and aspirated to dryness with a 28-gage needle. Subsequently, 30 μL of sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE) sample buffer (8 M urea, 2% sodium dodecyl sulfate, 100 mM dithiothreitol, 375 mM Tris, pH 6.8) was added to each sample. Immune complexes were dissociated by heating to 37º C for 2 h and resolved by SDS–PAGE. Gel electrophoresis was performed using 10% polyacrylamide gels. Proteins were transferred to PVDF membranes using a semi-dry transfer system and immunoblotted with either rabbit anti-A2AR polyclonal antibody (Ciruela et al. 2004) or rabbit anti-D2R polyclonal antibody (Millipore), and then horseradish peroxidase-conjugated goat anti-rabbit IgG (1/60000; Thermo Fisher Scientific, Inc., Rockford, IL, USA). The immunoreactive bands were developed using a chemiluminescent detection kit (Pierce, Rockford, IL, USA) (Ciruela and McIlhinney 1997).

BRET assay

For BRET experiments, HEK293 cells, transiently transfected with constant (1 μg) amount of plasmid encoding A2ARRluc and increasing amounts (0.5–6 μg) of plasmids encoding for D2RYFP or D2RmutYFP, were rapidly washed twice in phosphate-buffered saline, detached, and resuspended in the same buffer. Cell suspensions (20 μg protein) were distributed in duplicate into 96-well microplate black plates with a transparent bottom (Corning 3651; Corning, Stockholm, Sweden) for fluorescence measurement or white plates with a white bottom (Corning 3600) for BRET determination. For BRET1 measurement, h-coelenterazine substrate (Life Technologies Corp., Grand Island, NY, USA) was added at a final concentration of 5 μM, and readings were performed 1 min after using the POLARstar Optima plate-reader (BMG Labtech) that allows the sequential integration of the signals detected with two filter settings [485 nm (440–500 nm) and 530 nm (510–560 nm)]. The BRET ratio was defined as previously described (Canals et al., 2003; Ciruela et al. 2004; Woods and Ferre 2005; Woods et al. 2005).

SNAP labeling

Eventually, we identified cells to perform FRET experiments and also quantified the expression of the D2R constructs by means of the SNAP-tag technology (see above). Briefly, cells transfected with the D2RAGT and D2RmutAGT constructs were seeded into 18 mm diameter glass coverslips or in 96-wells plates and then stained with 5 μM or 100 μM, respectively, of the SNAP-Surface 647 ligand (New England BioLabs, Ipswich, MA, USA) in supplemented Dulbecco's modified Eagle's medium during 30 min at 37°C, 5% CO2. Finally, cells were washed three times with phenol red-free Hank's balanced salt solution containing 1 g/L glucose (HBSS: 137 mM NaCl, 5.4 mM KCl, 0.3 mM Na2HPO4, 0.4 mM KH2PO4, 4.2 mM NaHCO3, 1.3 mM CaCl2, 0.5 mM MgCl2, 0.6 mM MgSO4, 5.6 mM glucose, pH 7.4) and imaged at 688 nm upon excitation at 647 nm with a Cy5 filter set (Zeiss, Oberkochen, Germany) or fluorescence measured at 680 nm upon excitation at 640 nm in a POLARstar Optima plate-reader (BMG Labtech).

Microscopic FRET measurements

Binding of MRS5424 in cells expressing the A2ARCFP or PTHR1CFP construct was determined by real-time single-cell FRET experiments. In brief, transiently transfected cells seeded into 18 mm diameter glass coverslips were mounted in an Attofluor holder and placed on an inverted Axio Observer microscope (Zeiss) equipped with a 63× oil immersion objective and a dual-emission photometry system (TILL Photonics, Gräfelfing, Germany). Then, cells were continuously superfused with the fluorescent ligand dissolved in HBSS, applied with the aid of a focal drug application system (OCTAFLOW; ALA Scientific Instruments, Westbury, NY, USA). A Polychrome V (Till Photonics) was used as the light source and signals detected by avalanche photodiodes were digitized using a Digidata 1440A analog/digital converter (Molecular Devices, Sunnyvale, CA, USA). pCLAMP (Molecular Devices) and GraphPad Prism (GraphPad Software, La Jolla, CA, USA) softwares were used for data collection and analysis. FRET was measured upon excitation at 430 ± 10 nm (beam splitter dichroic long-pass 460 nm) and an illumination time set to 10 ms at 10 Hz. Then, the emission light intensities were determined at 535 ± 15 nm (F535) and 480 ± 20 nm (F480) with a beam splitter dichroic long-pass of 505 nm. No corrections for spillover between channels or direct Alexa532 excitation were made. The increase in FRET ratio (F535/F480) was fitted to the equation: r(t) = A × (1 − et), where τ is the time constant (s) and A is the magnitude of the FRET signal. When necessary for calculating τ, agonist-independent changes in FRET as a result of photobleaching were subtracted. Cell images were taken with a Zeiss AxioCamMR3 and processed with the Axiovision 4.8 software (Zeiss).

Statistics

The number of samples (n) in each experimental condition is indicated in the figure legends. When two experimental conditions were compared, statistical analysis was performed using an unpaired t-test. Otherwise, statistical analysis was performed by one-way analysis of variance (anova) followed by the Student–Newman–Keuls post hoc test. Statistical significance was set as p < 0.05.

Results

Role of the D2R IL3 Arg-rich domains on A2AR/D2R oligomerization

It is well established that a Coulombic interaction between the C-terminal tail of the A2AR and IL3 of the D2R is involved in the A2AR/D2R oligomerization (Ciruela et al. 2004). Interestingly, although the role of the A2AR C-terminal tail in this phenomenon has been largely studied (Borroto-Escuela et al. 2010a,b,a,b), the impact of Arg-rich domains of the D2R IL3 on the oligomeric function has been less explored. Thus, we aimed to investigate the role of two regions rich in positive charges within the D2R IL3 on the negative allosteric receptor–receptor interaction associated with the A2AR/D2R oligomer. To this end, and by means of site-directed mutagenesis, we generated a mutated D2R construct (D2RmutYFP), where the arginine (Arg) residues in IL3 of the D2R were substituted by alanine (Ala) (Fig. 1a). First, we examined whether the Arg to Ala substitutions affected the functionality of the receptor, as it had to be challenged in further experiments with the selective agonist quinpirole, and a loss of efficacy could lead to misinterpretation of the results. Therefore, by means of a luciferase reporter assay, we determined changes in CRE transcription, which is increased after the activation of the CREB. Specifically, we examined the ability of both D2RYFP and D2RmutYFP to antagonize the forskolin-mediated activation of AC through coupling to Gαo/i, as previously described (Obadiah et al. 1999). Interestingly, both D2R constructs equally reduced the forskolin-mediated cAMP accumulation after quinpirole (25 nM) challenge (Fig. 1b), suggesting that the Arg to Ala substitutions did not affect the receptor functionality.

Figure 1.

Characterization of the D2R construct. (a) Schematic representation of the D2Rmut construct. The mutations performed at the third intracellular loop 3 (IL3) are indicated. The D2R structure was suited using the PDB 1JGJ of rhodopsin and prepared using PyMOL (PyMOL Molecular Graphics System, DeLano Scientific, San Carlos, CA, USA). (b) Determination of cAMP accumulation by means of the luciferase reporter assay system. HEK293 cells transiently transfected with pGL4-CRE-luc2p/phRG-B plus D2RYFP (red bars) or D2RmutYFP (blue bars) were incubated either with forskolin (1 μM) plus Hank's balanced salt solution buffer (solid bars) or quinpirole (25 nM) (empty bars). Light emission was normalized assigning the 100% of effect to that obtained when incubating cells with forskolin. Data are expressed as the mean ± SEM of three independent experiments. (*) indicate statistically significant differences (p < 0.05; Student's t-test) when comparing Hank's balanced salt solution versus quinpirole treatment in both D2RYFP and the D2RmutYFP transfected cells.

Next, we evaluated the ability of the D2RmutYFP to oligomerize with the A2AR. To this end, we examined the physical interaction with the A2AR by means of a classical biochemical approach (i.e., co-immunoprecipitation experiments) and by a biophysical assay (i.e., BRET experiments). Thus, from extracts of HEK293 cells transiently co-transfected with A2ARCFP plus D2RYFP or D2RmutYFP, the mouse anti-A2AR antibody co-immunoprecipitated a protein of molecular weight ~75 kDa corresponding to the D2RYFP (either the wild type or the mutant) (Fig. 2a, IP: anti-A2AR, upper panel, lanes 4 and 5). Importantly, this protein band did not appear in singly transfected (A2ARCFP, D2RYFP or D2RmutYFP) cells immunoprecipitated with the same antibody (Fig. 2a, IP: anti-A2AR, upper panel, lanes 1, 2, and 3). Finally, we confirmed the presence of the A2AR by means of a rabbit polyclonal antibody (Fig. 2a, IP: anti-A2AR, lower panel, lanes 1, 4, and 5). Overall, these results suggested that by mutating the Arg residues of IL3 of the D2R, its interaction with the A2AR was not qualitatively altered.

Figure 2.

Role of the D2R third intracellular loop 3 Arg-rich domains on A2AR/D2R oligomerization. (a) Co-immunoprecipitation experiments. Membrane suspensions of HEK293 cells transiently expressing A2ARCFP (lane 1), D2RYFP (lane 2), D2RmutYFP (lane 3), A2ARCFP plus D2RYFP (lane 4), A2ARCFP plus D2RmutYFP (lane 5) were solubilized and processed for immunoprecipitation using mouse anti-A2AR monoclonal antibody (IP). Immunoprecipitates were analyzed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis and immunoblotted using rabbit anti-D2R polyclonal antibody and rabbit affinity-purified anti-A2AR polyclonal antibody (IB). The presented blots are representative of three different experiments with similar qualitative results. (b) BRET experiments. BRET saturation curves for the A2AR/D2R oligomers (A2ARRluc/D2RYFP: filled red circles and A2ARRluc/D2RmutYFP: filled blue circles) at increasing expression levels of the YFP-tagged vectors. Plotted on the X-axis is the fluorescence value obtained from the YFP, normalized with the luminescence value of A2ARRluc 10 min after h-coelenterazine incubation. Results are expressed as mean ± SEM (n = 8 in triplicate). (c) Comparison of the BRETmax obtained in (b). Values represent percentages of maximal saturable BRET responses (BRETmax). Mean ± SEM. (n = 8 in triplicate). ***: A2ARRluc/D2RmutYFP group is significantly different compared with the A2ARRluc/D2RYFP group (< 0.001); +++: control group (A2ARRluc+D2RmutYFP) is significantly different compared with the A2ARRluc/D2RmutYFP and A2ARRluc/D2RYFP groups (< 0.001).

Also, we analyzed the degree of the A2AR-D2R interaction by means of BRET experiments. Thus, a BRET saturation curve was constructed in HEK293 cells co-transfected with a constant amount of an A2ARRluc construct, while increasing the concentrations of plasmids containing D2RYFP or D2RmutYFP. Interestingly, a positive BRET signal was obtained for the transfer of energy between A2ARRluc and both the D2RYFP and the D2RmutYFP. The BRET signal, as reflected in the BRET ratio, increased as a hyperbolic function of the concentration of the D2RYFP fusion constructs (Fig. 2b). However, the pairing of A2ARRluc and D2RmutYFP led to a substantial reduction in the maximal BRET signal (BRETmax) when compared with the A2ARRluc and D2RYFP pair (45 ± 2 mBU and 81 ± 2 mBU, respectively) (Fig. 2b). Importantly, when cells singly expressing the A2ARRluc and D2RYFP-expressing cells were mixed, no BRET signal was observed (Fig. 2c). Overall, the results obtained suggested that while the Arg to Ala substitutions within the D2R IL3 did not abolish A2AR/D2R oligomerization, the proximity between the two receptors was significantly reduced. Alternatively, it could also be possible that the mutations would lead to a lower proportion of oligomers, thus the partial BRET signal detected would be the sum of a smaller number of direct receptor–receptor interactions.

D2R-mediated allosteric modulation of A2AR agonist binding

Antagonistic A2AR-mediated allosteric modulation of D2R function has been largely studied; however, the reverse condition, that is, D2R-mediated allosteric modulation of the A2AR, is still elusive. Consequently, we decided to investigate both the possible allosteric modulation exerted by the D2R on A2AR function, when these receptors are forming part of an oligomer, and also, the contribution of Arg-rich domains of IL3 of the D2R to this receptor–receptor allosterism. To this end, we implemented a real-time FRET-based A2AR agonist binding procedure in cells co-expressing both A2ARs and D2Rs. Accordingly, we recorded the FRET engaged between a novel fluorescent A2AR agonist synthesized in this study (MRS5424) and the A2ARCFP construct (Fig. 3b). First, we checked that the agonist-receptor interaction was observed at the cell surface of the transfected cells, and that the binding was completely blocked in the presence of an A2AR-specific antagonist, ZM241,385 (Fig. 3a). And thereafter, a specific FRET signal between MRS5424 and the A2ARCFP was measured in a real-time mode (Fig. 3c). Thus, under these experimental conditions, we analyzed changes in the FRET signal exerted by a quinpirole challenge on cells expressing A2ARCFP plus either D2R or D2Rmut. It is important to mention here that to ensure the presence of both A2AR and D2R within the same assayed cell, we engineered a D2RAGT construct (Fig. 4a), which allowed the subsequent visualization of the receptor at the cell surface (Fig. 4b). Indeed, in Fig. 4b, it is possible to visualize that the A2ARCFP and the D2R-AGT fusion constructs (D2RAGT and D2RmutAGT) labeled with the SNAP substrate are co-distributed at the plasma membrane of co-transfected cells, thus providing the ideal conditions to assess our real-time FRET-based A2AR agonist binding experiments.

Figure 3.

Determination of A2AR agonist binding by real-time fluorescence resonance energy transfer (FRET) in single living cells. (a) Specific MRS5424 binding to the A2ARCFP. HEK293 cells transiently transfected with A2ARCFP were superfused with MRS5424 (2 μM) during 5 min, washed and observed on an inverted microscope without (upper image) or with (down image) ZM241,385 (1 μM), a specific A2AR antagonist. Scale bar: 10 μm. (b) Schematic representation of the FRET experiment between the fluorescent A2AR agonist MRS5424 (APECAlexa532) and the A2ARCFP construct. The A2AR and CFP structures were prepared using PyMOL and the PDBs 3EML and 1EMA, respectively. (c) Time-resolved changes in CFP and Alexa532 fluorescence emission signals in single cells transfected with A2ARCFP. The emission intensities of CFP (F480, blue trace), Alexa 532 (F535, yellow trace), and the ratio F535/F480 (black trace) in response to MRS5424 were recorded simultaneously from single HEK293 cells expressing the A2ARCFP. Shown are the changes induced by rapid superfusion with 2 μM MRS5424. The increase of the ratio F535/F480 was fitted by a simple monoexponential curve giving a time constant (τ) in this experiment of 12.0 ± 0.3 s. Traces are representative of eight separate experiments.

Figure 4.

Determination of A2AR agonist binding by real-time fluorescence resonance energy transfer in single living cells. (a) Schematic representation of the A2ARCFP and D2RAGT constructs. The schematic alkyltransferase (AGT) (O6-alkylguanine-DNA alkyltransferase) diagram was prepared using PyMOL and PDB 1EH6. (b) Cell surface localization of the A2ARCFP and D2RAGT constructs in living cells. HEK293 cells were transiently transfected with A2ARCFP and either D2RAGT or D2RmutAGT, stained with the SNAP substrate and visualized at 480 nm (CFP) and 688 nm (SNAP) upon excitation at 430 nm and 647 nm, respectively, using an inverted microscope. Superposition of images (merge) revealed A2ARCFP and D2RAGT co-distribution at the cell surface. Scale bar: 10 μm. (c) Cell surface quantification of the A2ARCFP and D2RAGT constructs in living cells. HEK293 cells were transiently transfected with A2ARCFP and either D2RAGT or D2RmutAGT, stained with the SNAP substrate and plated in wells, where fluorescence at 680 nm was measured upon excitation at 640 nm using a POLARstar Optima plate-reader. Results are expressed as mean ± SEM (n = 3 in triplicate), after the subtraction of the fluorescence measured in mock cells (AU, arbitrary units).

First, we recorded FRET in cells singly transfected with the A2ARCFP. Thus, upon excitation at 430 nm (CFP excitation), the signals recorded from single HEK293 cells expressing A2ARCFP challenged with the fluorescent agonist were analyzed at emissions of 480 nm (Fig. 3b, lower panel, CFP) and 535 nm (Fig. 3b, lower panel, Alexa532), and the corresponding normalized ratio F535/F480 plotted (Fig. 3b, upper panel). After addition of MRS5424 (2 μM), a symmetrical decrease in CFP emission and an increase in Alexa532 emission indicated that the change was indeed because of an increase in FRET between A2ARCFP and MRS5424 as a consequence of the ligand binding (i.e., increase in the ratio F535/F480) (Fig. 3b). Interestingly, control experiments with cells expressing PTHRCFP showed no FRET when MRS5424 was superfused on these cells (data not shown). On the other hand, the selection of the MRS5424 concentration (2 μM) was based on previous results (Fernández-Dueñas, unpublished data), indicating that this concentration elicited the best-defined response to be subsequently modulated with the D2R agonist.

Next, we analyzed the D2R-mediated allosteric modulation of A2AR agonist binding in cells co-expressing the A2ARCFP and D2RAGT constructs. Interestingly, the proteins were shown to co-localize at the cell surface (Fig. 4b), and it could also be assessed that the plasma membrane expression of the D2RAGT constructs were unaffected because of the mutations (Fig. 4c). Thus, upon identification of a doubly transfected cell (as in Fig. 4b), we proceeded to analyze the binding of MRS5424 at the A2ARCFP in the absence or the presence of the D2R agonist quinpirole (100 μM). Importantly, MRS5424 binding was unaffected by quinpirole in cells transfected only with the A2ARCFP (data not shown). Thus, upon D2RAGT co-expression and quinpirole challenge, a significantly lower FRET signal was observed, indicating that D2R activation diminished the binding of the A2AR agonist (Fig. 5a, left panel). Noteworthy, in this Figure, the inverse of the decline of CFP fluorescence (1/FCFP) was used as readout of the ligand binding event, because the diminution of CFP fluorescence was exclusively as a result of FRET and also to avoid any contamination of the FRET determinations. Furthermore, we analyzed the kinetic data by fitting the CFP decline to a monoexponential decay curve, and a decrease was observed in the association rate upon addition of quinpirole (0.134 ± 0.025 s−1 with quinpirole vs. 0.028 ± 0.005 s−1 with buffer). This indicated that the association of the fluorescent ligand was significantly affected by the conformational change of the D2R upon quinpirole activation. Conversely, in cells co-expressing the A2ARCFP and D2RmutAGT, the binding of the fluorescent A2AR ligand was unaffected by quinpirole superfusion (Fig. 5a, right panel). In addition, when adjusting the CFP decline to a monoexponential decay curve, the kinetic data revealed that the association rate constant was not as affected by the D2RmutAGT (0.046 ± 0.007 s−1) as with the wild-type D2RAGT. Therefore, Arg mutations diminished the ability of the D2R to exert a complete allosteric modulation of binding at the A2AR. Overall, these results demonstrated that the IL3 Arg-rich domains of D2R played a major role in the allosteric effects of the D2R within the A2AR/D2R oligomer, as the negative allosteric modulation exerted by the D2R (74.3 ± 3.1%) disappeared (94.4 ± 4.3%; p < 0.05) following mutagenesis (Fig. 5b).

Figure 5.

Real-time fluorescence resonance energy transfer (FRET) determinations of the D2R-mediated modulation of A2AR agonist binding. (a) Time-resolved changes in FRET signals (see Fig. 3) in cells transfected with A2ARCFP plus D2RAGT or A2ARCFP plus D2RmutAGT in the absence (black trace) or presence (red and blue traces) of quinpirole (100 μM) were recorded. The FRET increase (1/F480) was fitted by a simple monoexponential curve and the magnitude of the FRET signal (A, see Materials and methods) calculated for each experimental condition. Traces are representative of five separate experiments with similar qualitative and quantitative results. (b) The variation of the A value for each experimental condition was plotted. Data represent the average ± SEM values of five independent experiments. Asterisk indicates data significantly different from the control condition (i.e., in the absence of quinpirole): *p < 0.05 by anova with Student–Newman–Keuls multiple comparison post hoc test.

Functional consequences of the D2R allosteric modulation

In view of the previous results, one would expect that D2R-mediated allosteric modulation of the A2AR would lead to functional consequences on A2AR signaling. Therefore, we evaluated the effects of the D2RAGT or the D2RmutAGT challenge on A2AR-mediated AC stimulation by measuring cAMP accumulation in co-transfected cells. It is important to mention here that in cells singly transfected with A2ARCFP the selective A2AR agonist CGS21680 (50 nM) led to an increase of cAMP accumulation, and while the A2AR antagonist ZM241385 (1 μM) totally blocked this effect, the D2R agonist quinpirole (25 nM) did not produce any effect on A2AR-mediated cAMP accumulation (data not shown). Interestingly, in doubly A2ARCFP-D2RAGT transfected cells quinpirole partially and significantly (~60%; p < 0.01) diminished the effects of CGS21680 on cAMP accumulation (Fig. 6a, red bars, CGS+Quin). On the other hand, when cells expressing A2ARCFP and D2RmutAGT were challenged with quinpirole, a significant (~80%; p < 0.01) reduction of A2AR agonist-mediated cAMP accumulation was observed (Fig. 6a, red bars, CGS+Quin). Curiously, when inhibition of D2RAGT- and D2RmutAGT-mediated cAMP accumulation was compared, it was shown to be significantly (p < 0.05) different (Fig. 6a). Apparently, these contradictory results might be readily explained if we consider that the establishment of an A2AR/D2R functional oligomer (i.e., displaying receptor–receptor allosterism) would allow direct A2AR–D2R cross talk, providing a balanced outcome of Gαs- and Gαi -mediated AC modulation (Fig. 6b). On the contrary, when no direct functional A2AR-D2R cross talk would exist, a shift to more Gαi-coupled D2Rs would happen, thus leading to an increased AC inhibition outcome (Fig. 6b).

Figure 6.

Functional consequences of the D2R allosteric modulation. (a) Allosteric modulation of the A2ARCFP activity. HEK293 cells transiently transfected with pGL4-CRE-luc2p/phRG-B and A2ARCFP plus D2RAGT (red bars) or D2RmutAGT (blue bars) were incubated with forskolin (1 μM), CGS21680 (50 nM), CGS + ZM241385 (50 nM + 1 μM, respectively), or CGS21680 + quinpirole (50 nM + 25 nM, respectively) and the cAMP accumulation determined by means of the luciferase reporter assay system (see Materials and methods). Light emission was normalized assigning the 100% of effect to that obtained when incubating cells with forskolin. Data are expressed as the mean ± SEM of three independent experiments. (*) indicates statistically significant differences (< 0.05; Student's t-test) when comparing cells transfected with the D2RAGT or the D2RmutAGT. (b) Schematic representation of the oligomeric antagonistic A2AR–D2R interaction. In the left part, the native A2AR/D2R oligomer is shown, in which a balance between Gαs- and Gαi/o-coupling would exist, and the bidirectional negative allosteric receptor–receptor interaction may outline the final adenylyl cyclase (AC) signaling efficacy. Indeed, at the G-protein level the expected antagonistic interaction can be observed as the A2AR protomer via Gαs activates AC, while the D2R protomer would partially lose its Gαi/o-coupled AC inhibition. In the right part, the mutated A2AR/D2R oligomer is shown. This oligomer lacks the negative allosteric receptor–receptor interaction and non-A2AR–D2R trans-inhibition phenomena are established. Thus, in the absence of the powerful antagonistic A2AR–D2R interaction at the level of D2R recognition and Gαi/o-coupling, an enhanced AC inhibition is observed.

Overall, the results of this study show that the Arg-rich domains of the D2R IL3 played a key functional role in the A2AR/D2R oligomer. Thus, when mutating these residues it could be observed that although (i) there were no changes on membrane density of D2R and (ii) no effect on D2R signaling (iii) it was affected but not abolished A2AR-D2R heteromerization, and (iv) a selective abolition of the allosteric interactions between A2AR and D2R occurred.

Discussion

The molecular interactions between the A2AR and the D2R when engaged in an oligomeric complex have been postulated to play a major role in the development and also the treatment of neurodegenerative disorders, such as Parkinson's disease (for review see (Fuxe et al. 2010)). As a result, the elucidation of the mechanisms mediating such receptor–receptor interactions has become a main goal during recent years. Consequently, as commented earlier, it was previously shown that a Coulombic interaction established between the IL3 of the D2R and the C-terminal tail of the A2AR plays a pivotal role in the phenomenon of A2AR/D2R oligomerization (Canals et al., 2003; Ciruela et al. 2004; Woods and Ferre 2005; Woods et al. 2005). Thus, in the present work, we aimed to examine the allosteric modulation that the D2R exerts on the A2AR within the A2AR/D2R oligomer. In addition, we also aimed to demonstrate that this receptor–receptor interaction is mostly mediated by the two highly positively charged regions located at the IL3 of the D2R, not only at the level of the binding of A2AR agonists but also with respect to the functionality of the latter receptor.

Hence, we performed site-directed mutagenesis to generate a D2R mutant in which the positively charged Arg residues were replaced by uncharged alanine residues. Interestingly, the mutation of the Arg residues did not affect the functionality of the receptor, as it was able to inhibit forskolin-induced cAMP accumulation at the same level as the wild-type D2R. Accordingly, it could be assumed that the results obtained when examining the allosteric effects of the D2R agonist were not because of an aberrant signaling pathway as a result of the mutations. Interestingly, by means of co-immunoprecipitation experiments, we found that the Arg-rich domains of IL3 of the D2R seemed non-essential for A2AR and D2R oligomerization as D2Rmut still co-immunoprecipitated with the A2AR. However, the Arg→Ala substitutions within the D2R IL3 probably reduced the molecular proximity between these two receptors as indicated in our BRET assays. These results are consistent with those previously obtained concerning the role of negatively charged residues on the A2AR C-terminal tail, which in fact may interact electrostatically with IL3 of the D2R in A2AR/D2R oligomerization (Borroto-Escuela et al. 2010a,b). Although these Coulombic interactions disappeared, the complex was still formed, as the protein–-protein interaction also depends on other receptor regions, for instance the TMs-IV and V of the D2R (Borroto-Escuela et al. 2010a,b). Needless to say, it could also be possible for the receptor–receptor interaction to be wholly affected and that we were observing a lower proportion of oligomers formed. However, when assessing the expression levels of the D2RAGT constructs, we did not find differences in the presence of the wild type and the mutant receptor at the cell surface. Nevertheless, it could be concluded that the mutations affected in some manner the present receptor–receptor interaction, thus the possible effects on the presumably antagonistic allosteric interaction were subsequently examined.

Interestingly, regarding the allosteric modulation of the D2R on A2AR binding, the generation of the mutant variant of the D2R permitted to demonstrate the relevance of the positively charged residues located at the IL3 in such receptor–receptor interaction. Thus, upon co-transfecting both the D2RmutAGT and the A2ARCFP, quinpirole treatment did not affect FRET between the fluorescent A2AR agonist and A2ARCFP, as compared with the wild-type D2R. Previous studies have revealed that an intramembrane interaction occurs between the A2AR and the D2R; thus, it was shown that the A2AR inhibits the D2R-mediated neuronal excitability and neurotransmitter release and that the D2R negatively modulates A2AR ligand binding to inhibit A2AR-mediated protein phosphorylation and gene expression (Canals et al., 2003; Ciruela et al. 2004; Woods and Ferre 2005; Woods et al. 2005). However, to our knowledge, this is the first time the negative allosteric modulation of the D2R on A2AR agonist binding has been visualized in a real-time mode. Thus, we were able to determine that D2R activation partially inhibited and also slowed the binding association of the fluorescent A2AR agonist. Furthermore, the results obtained clearly indicated that this important interaction was abolished by mutating the IL3 of the D2R. In fact, we also evaluated the allosteric modulation of the D2R on A2AR agonist-mediated cAMP accumulation, and it was shown that the D2R agonist quinpirole, by activating the D2Rmut, was able to completely counteract A2AR agonist-mediated activation. Interestingly, it has been previously shown that the antagonistic allosteric A2AR/D2R interaction favors a rapid onset of β-arrestin-2 D2R signaling through Gαi/o-uncoupling (Canals et al., 2003; Ciruela et al. 2004; Trincavelli et al. 2012; Woods and Ferre 2005; Woods et al. 2005). Thus, when the receptor–receptor allosteric interaction disappears, as in the absence of Arg-rich domains of the D2R IL3, the D2R would be able to mostly signal through Gαi/o-protein to inhibit cAMP accumulation. Consequently, it could be considered that in the absence of A2AR–D2R allosteric modulation, a major outcome of Gαi/o-signaling depending cascades occurs (Fig. 6b). Interestingly, this condition might be exacerbated by the fact that the levels of Gαi/o-protein may be higher than the Gαs (Borroto-Escuela et al. 2011), thus contributing to the enhanced inhibition of AC activity. In this regard, it is currently thought that upon agonist stimulation and the subsequent receptor conformational change, this information is transmitted into the IL3 (Li et al. 1996) Thus, it would seem likely that changing some residues (Arg→Ala) of this domain would disrupt its normal function. Consequently, the lack of an allosteric modulation of the D2Rmut could also be explained, not only because a reduced direct receptor–receptor interaction within the A2AR/D2R oligomer but also by means of other mechanisms located at the intracellular signal transmission pathways (Kull et al. 1999; Zurn et al. 2009). However, our results point to a scenario in which the mutation of the Arg residues at the IL3 reduced the physical interaction between the D2R and the A2AR, and therefore partially precluded the D2R-mediated allosteric modulation of the A2AR.

In conclusion, the findings of this study provide further evidence of the existence of antagonistic A2AR-D2R interactions. In particular, the Arg residues (217–222 and 267–269) on IL3 of the D2R have been identified to play a major key role in such receptor–receptor allosteric interaction. Thus, it would be feasible to target this region in order to allosterically modulate the functionality of the A2AR/D2R oligomer.

Acknowledgements

This work was supported by grants SAF2011-24779 and Consolider-Ingenio CSD2008-00005 from Ministerio de Ciencia e Innovación and ICREA Academia-2010 from the Catalan Institution for Research and Advanced Studies to FC. Also, VF-D, MG-S, and FC belong to the “Neuropharmacology and Pain” accredited research group (Generalitat de Catalunya, 2009 SGR 232). Support to KAJ and TSK from the NIDDK Intramural Research Program of the National Institutes of Health, Bethesda, MD, USA is acknowledged. We also thank Esther Castaño, Eva Julià, and Benjamín Torrejón, from the Scientific and Technical Services (SCT)-Bellvitge Campus of the University of Barcelona for the technical assistance.

Conflict of interest

The authors declare no conflict of interest.

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