Original Article

Molecular Psychiatry (2012) 17, 650–662; doi:10.1038/mp.2011.93; published online 16 August 2011

Dopamine D4 receptor, but not the ADHD-associated D4.7 variant, forms functional heteromers with the dopamine D2S receptor in the brain

S González1, C Rangel-Barajas2, M Peper3, R Lorenzo3, E Moreno1, F Ciruela4, J Borycz5, J Ortiz6, C Lluís1, R Franco7, P J McCormick1, N D Volkow8, M Rubinstein3, B Floran2 and S Ferré5

  1. 1Department of Biochemistry and Molecular Biology, Centro de Investigación Biomédica en Red sobre Enfermedades Neurodegenerativas, University of Barcelona, Barcelona, Spain
  2. 2Departamento de Fisiología, Biofísica y Neurociencias, Centro de Investigación y de Estudios Avanzados del Instituto Politécnico Nacional, México D.F., México
  3. 3Departamento de Fisiología y Biología Molecular y Celular, Instituto de Investigaciones en Ingeniería Genética y Biología Molecular, Consejo Nacional de Investigaciones Científicas y Técnicas, Buenos Aires, Argentina
  4. 4Unitat de Farmacologia, Departament Patologia i Terapèutica Experimental, Universitat de Barcelona, L’Hospitalet de Llobregat, Barcelona, Spain
  5. 5CNS Receptor-Receptor Interactions Unit, National Institute on Drug Abuse, Intramural Research Program, National Institutes of Health, Baltimore, MD, USA
  6. 6Department of Biochemistry and Molecular Biology, Neuroscience Institute, Universitat Autónoma de Barcelona, Bellaterra, Spain
  7. 7Departamento de Neurociencias, Centro de Investigación Médica Aplicada, Universidad de Navarra, Pamplona, Spain
  8. 8National Institute on Drug Abuse, National Institutes of Health, Bethesda MD, USA

Correspondence: Dr S Ferré, National Institute on Drug Abuse, Intramural Research Program, National Institutes of Health, 251 Bayview Boulevard, Baltimore, MD 21224, USA. E-mail: sferre@intra.nida.nih.gov

Received 25 March 2011; Revised 27 June 2011; Accepted 7 July 2011
Advance online publication 16 August 2011

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Abstract

Polymorphic variants of the dopamine D4 receptor have been consistently associated with attention-deficit hyperactivity disorder (ADHD). However, the functional significance of the risk polymorphism (variable number of tandem repeats in exon 3) is still unclear. Here, we show that whereas the most frequent 4-repeat (D4.4) and the 2-repeat (D4.2) variants form functional heteromers with the short isoform of the dopamine D2 receptor (D2S), the 7-repeat risk allele (D4.7) does not. D2 receptor activation in the D2S–D4 receptor heteromer potentiates D4 receptor-mediated MAPK signaling in transfected cells and in the striatum, which did not occur in cells expressing D4.7 or in the striatum of knockin mutant mice carrying the 7 repeats of the human D4.7 in the third intracellular loop of the D4 receptor. In the striatum, D4 receptors are localized in corticostriatal glutamatergic terminals, where they selectively modulate glutamatergic neurotransmission by interacting with D2S receptors. This interaction shows the same qualitative characteristics than the D2S–D4 receptor heteromer-mediated mitogen-activated protein kinase (MAPK) signaling and D2S receptor activation potentiates D4 receptor-mediated inhibition of striatal glutamate release. It is therefore postulated that dysfunctional D2S–D4.7 heteromers may impair presynaptic dopaminergic control of corticostriatal glutamatergic neurotransmission and explain functional deficits associated with ADHD.

Keywords:

dopamine receptors; receptor heteromers; ADHD; striatum; glutamate

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Introduction

Dopamine D4 receptors are expressed in the prefrontal cortex, in GABAergic interneurons and in glutamatergic pyramidal neurons, including their striatal projections.1, 2, 3 D4 receptors have been implicated in attention-deficit hyperactivity disorder (ADHD).1, 4, 5, 6 In fact, the prefrontal cortex and associated fronto-striatal circuits are critical for executive function and are involved in ADHD.5 The gene encoding the human D4 receptor contains a large number of polymorphisms in its coding sequence.4 The most extensive polymorphism is found in exon 3, a region that codes for the third intracellular loop (3IL) of the receptor. This polymorphism consists of a variable number of tandem repeats in which a 48-bp sequence exists as 2- to 11-fold repeats.7 The three most common variants contain 2, 4 and 7 repeats (D4.2, D4.4 and D4.7, respectively). D4.4 constitutes the most frequent variant, with a global frequency of 64%, followed by D4.7 (21%) and D4.2 (8%).8 Importantly, a high prevalence of the D4.7 variant has been demonstrated in children diagnosed with ADHD.5 Though stimulation of the D4,7 variant has been reported to be less potent at inhibiting cAMP than D4.2 or D4.4,9 the functional significance of these variants are poorly understood.

Receptor heteromers are becoming the focus of extensive research in the field of G-protein-coupled receptors.10 A receptor heteromer is currently defined as a macromolecular complex composed of at least two (functional) receptor units with biochemical properties that are demonstrably different from those of its individual components.10 In some cases, receptor heteromers provide a framework in which to understand the role of receptors with no clear functional significance, and example being the D3 receptor, which forms heteromers with the D1 receptor and modifies its function.11 A recent study showed that in mammalian transfected cells, the long isoform of the D2 receptor (D2L) heteromerizes with the three main D4 receptor variants, D4.2, D4.4 and D4.7.12 Interestingly, results from the same study suggested that D4.7 was less effective in forming heteromers with D2L receptors.12 In view of the reported evidence of predominant co-localization of D4 receptors with the short isoform of the D2 receptor (D2S) in corticostriatal glutamatergic terminals,2, 3, 13 we first investigated if any of the three main human variants of the D4 receptor could interact both physically and functionally with D2S. By using the Bioluminescence Resonance Energy Transfer (BRET) technique, here we show evidence for the formation of heteromers between D2S and D4.2 and D4.4 variants of the D4 receptor. In contrast, the D4.7 variant failed to form heteromers with the D2S receptor. In transfected cells, we found a biochemical property of the D2S–D4 receptor heteromer, which consists of the ability of D2S receptor activation to potentiate D4 receptor-mediated mitogen-activated protein kinase (MAPK) signaling. A similar result was observed in striata from wild-type (WT) mice, a species that expresses D4 receptors with a short 3IL comparable to human D4.2. In contrast, potentiation of D4 receptor-mediated MAPK signaling was not observed in transfected cells expressing D4.7 or in striata taken from knockin mice carrying a humanized 7-repeat intracellular loop identical to that found in human D4.7. Finally, analyzing neurotransmitter release in striatal slices and with in vivo microdialysis in rats, evidence was obtained for a key role of D2–D4 receptor interaction in the modulation of striatal glutamatergic neurotransmission.

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Materials and methods

Fusion proteins and expression vectors

The synthetic cDNAs for the human D4.2, D4.4 and, D4.7 receptor gene (kindly provided by TP Sakmar, Rockefeller University, USA) were amplified using sense oligonucleotide primer (5′-TCAACGGGACTTTCCAAAATGT-3′) and antisense primer (5′-CTCCGAGATCAACTTCTGCTCGCTTCGGTTACCC-3′), resulting in a cDNA fragment of 200bp. A second product was generated using the sense oligonucleotide primer (5′-AAGTTGATCTCGGAGGAAGATACAGCAGATGCAG-3′) and antisense primer (5′- GCGAATTCGCAGCAAGCACGTAGAGCCTTACG-3′), resulting in a cDNA fragment of 1500bp. Equimolar quantities of both fragments were used to produce a third product corresponding to the myc-D4.2, myc-D4.4 or myc-D4.7-tagged gene using the sense primer (5′-GTGCTCGAGCACCATGGGTAACCGAAGCACAG-3′) and antisense primer without its stop codon (5′-GCGAATTCTCAGCAGCAAGCACGTAGAGCCTTACG-3′), harboring unique XhoI and EcoRI restriction sites, respectively. The fragments were then subcloned in-frame into XhoI/EcoRI sites of the pcDNA3.1 vector (Invitrogen, Paisley, Scotland, UK). Next, the human cDNAs for the adenosine A1 receptor and dopamine D4.2, D4.4, D4.7 and D2S receptors, cloned in pcDNA3.1 were amplified without their stop codons using sense and antisense primers harboring unique XhoI and EcoRI sites to clone A1, D4.2, D4.4 and D4.7 receptors in the RLuc and the yellow fluorescent protein (YFP) corresponding vectors, and HindIII and BamHI to clone D2S in the RLuc and the YFP corresponding vectors. The mouse cDNAs for the D4 and D2S receptors, cloned in pCMV-SPORT6 (American Type Culture Collection, Manassas, USA) and pReceiver-M16 vectors, respectively (GeneCopoeia, Rockville, MD, USA), were amplified without their stop codons using sense and antisense primers harboring unique XhoI and EcoRV sites to clone D4 receptor in the RLuc corresponding vector, and XhoI and KpnI to clone D2S receptor in the RLuc and the YFP corresponding vectors. The amplified fragments were subcloned to be in-frame into restriction sites of the multiple cloning sites of EYFP-N3 vector (enhanced yellow variant of YFP; Clontech, Heidelberg, Germany) or the mammalian humanized pRluc-N1 vectors (Perkin-Elmer, Waltham, MA, USA) to give the plasmids that express the receptors fused to either Rluc or YFP on the C-terminal end of the receptor (D4.2-RLuc, D4.4-RLuc, D4.7-Rluc, D2S-RLuc and A1-RLuc or D2S-YFP, D4.7-YFP and D1-YFP, respectively). All constructs were verified by nucleotide sequencing and the fusion proteins are functional and expressed at the membrane level (see Results).

Cell culture and transient transfection

HEK (human embryonic kidney)-293T cells were grown in DMEM (Dulbecco's modified Eagle's medium) (Gibco Paisley, Scotland, UK) supplemented with 2mM L-glutamine, 100Uml−1 penicillin/streptomycin and 5% (v/v) heat-inactivated fetal bovine serum (all supplements were from Invitrogen). CHO cell lines were maintained in á-MEM medium without nucleosides, containing 10% fetal calf serum, 50μgml−1 penicillin, 50μgml−1 streptomycin and 2mM L-glutamine (300μgml−1). Cells were maintained at 37°C in an atmosphere of 5% CO2, and were passaged when they were 80–90% confluent, twice a week. HEK-293T or CHO cells growing in six-well dishes or in 25cm2 flasks were transiently transfected with the corresponding fusion protein cDNA by the PEI (PolyEthylenImine; Sigma, Steinheim, Germany) method as previously described.14

Immunostaining

For immunocytochemistry, HEK-293T cells were grown on glass coverslips and transiently transfected with 1μg of cDNA corresponding to human D4.2-RLuc, D4.4-RLuc or D4.7-RLuc and 0.5μg of cDNA corresponding to human D2S-YFP or 0.8μg of cDNA corresponding to mouse D4-RLuc and 0.5μg of cDNA corresponding to mouse D2S-YFP. After 48h of transfection, cells were fixed in 4% paraformaldehyde for 15min and washed with phosphate-buffered saline contraining 20mM glycine to quench the aldehyde groups. After permeabilization with phosphate-buffered saline containing 0.05% Triton X-100 for 15min, cells were treated with phosphate-buffered saline containing 1% bovine serum albumin. After 1h at room temperature, cells were labeled with the primary rabbit monoclonal anti-human D4 receptor (1/10000; Abcam, Cambridge, UK) or with the primary goat polyclonal anti-D4 receptor (1/500; Santa Cruz Biotechnology, Santa Cruz, CA, USA) for 1h, washed and stained with the secondary antibody Cy3 anti-rabbit (1/200; Jackson ImmunoResearch, Baltimore, PA, USA) or with the secondary antibody Cy3 anti-goat (1/200; Jackson ImmunoResearch). The D2S-YFP construct was detected by its fluorescence properties. Samples were rinsed and observed in an Olympus confocal microscope.

BRET assay

HEK-293T cells were co-transfected with a constant amount of cDNA encoding for the receptor fused to RLuc and with increasingly amounts of cDNA encoding to the receptor fused to YFP to measure BRET as previously described.14 Both fluorescence and luminescence for each sample were measured before every experiment to confirm similar donor expressions (~100000 bioluminescence units) while monitoring the increase in acceptor expression (2000–20000 fluorescence units). The relative amounts of BRET acceptor are expressed as the ratio between the net fluorescence of the acceptor and the luciferase activity of the donor being the net fluorescence the fluorescence of the acceptor minus the fluorescence detected in cells only expressing the donor. The BRET ratio is defined as [(emission at 510–590)/(emission at 440–500)]−Cf, where Cf corresponds to (emission at 510–590)/(emission at 440–500) for the D4-RLuc or D2S-RLuc constructs expressed alone in the same experimental conditions. Curves were fitted by using a non-linear regression equation, assuming a single phase with GraphPad Prism software (San Diego, CA, USA).

Generation of knockin mutant mice carrying human expansions in the 3IL of the D4 receptor

A targeting vector was designed such that coding sequences of the 3IL of mouse Drd4 were replaced by human ortholog sequences corresponding to the most frequent 7-variable number of tandem repeat human variant allele (see Figure 4). The vector included a selectable PGK-neo cassette, flanked by two loxP sites, placed just downstream of Drd4 polyadenylation site and an herpes simplex virus-thymidine kinase cassette placed at one of the extremes of the targeting vector to select for the absence of random integrations. A long and short arm of Drd4 homology were inserted flanking the swapped sequence and the selectable marker, respectively. The linearized vector was used to electroporate hybrid 129svev/C57BL/6 ES cells (inGenious Targeting Laboratory, Stony Brook, NY, USA) and homologous recombinant clones were selected in the presence of G418 and gancyclovir. Two selected clones carrying the human 7-variable number of tandem repeat were used to microinject C57BL/6J blastocysts and one high percentage chimeric male mouse was used to produce heterozygote Drd4+/7repeat.neo mice. The neo cassette was excissed from the recombinant allele by crossing mutant mice with transgenic mice expressing Cre recombinase from an EIIa promoter (Jackson Laboratories; Cat. No. 003724). The resulting heterozygote Drd4+/7repeat (D4.7 knockin) mice were successively bred to C57BL/6J mice to obtain a congenic heterozygote strain (n=10) that was used to establish a breeding colony. Homozygous D4.7 knockin mice and their WT littermates were used for the experiments. Knockin animals were characterized as indicated in Figure 4.

Mouse striatal slices preparation

Mice were housed five per cage in a temperature (21±1°C) and humidity-controlled (55±10%) room with a 12:12-h light/dark cycle (light between 0800 and 2000hours) with food and water ad libitum. All animal procedures were conducted according to the standard ethical guidelines (National Institutes of Health Animal care guidelines and European Communities Council Directive 86/609/EEC) and approved by the Local Ethical and Animal Care Committees. Transgenic mice and littermattes were decapitated with a guillotine and the brains were rapidly removed and placed in ice-cold oxygenated (O2/CO2:95%/5%) Krebs-HCO3 buffer (124mM NaCl, 4mM KCl, 1.25mM NaH2PO4, 1.5mM MgCl2, 1.5mM CaCl2, 10mM glucose and 26mM NaHCO3, pH 7.4). The brains were sliced at 4°C in a brain matrix (Zivic Instruments, Pittsburgh, PA, USA) into 0.5mm coronal slices. Slices were kept at 4°C in Krebs-HCO3 buffer during the dissection of the striatum. Each slice was transferred into an incubation tube containing 1ml of ice-cold Krebs-HCO3 buffer. The temperature was raised to 23°C and after 30min, the media was replaced by 2ml Krebs-HCO3 buffer (23°C).

ERK phosphorylation assay

Striatal slices from transgenic mice and littermattes were incubated under constant oxygenation (O2/CO2:95%/5%) at 30°C for 4–5h in an Eppendorf Thermomixer (5 Prime, Boulder, CO, USA) with Krebs-HCO3 buffer. The media was replaced by 200μl of fresh Krebs-HCO3 buffer and incubated for 30min before the addition of ligands. Transfected CHO cells were cultured in serum-free medium for 16h before the addition of the indicated concentration of ligands for the indicated time. Both, cells and slices were lysed in ice-cold lysis buffer (50mM Tris-HCl pH 7.4, 50mM NaF, 150mM NaCl, 45mM β-glycerophosphate, 1% Triton X-100, 20μM phenyl-arsine oxide, 0.4mM NaVO4 and protease inhibitor cocktail). Cellular debris was removed by centrifugation at 13000g for 5min at 4°C and protein was quantified by the bicinchoninic acid method using bovine serum albumin dilutions as standard. To determine the level of extracellular signal-regulated kinases 1 and 2 (ERK1/2) phosphorylation, equivalent amounts of protein (10μg) were separated by electrophoresis on a denaturing 10% sodium dodecyl sulfate-polyacrylamide gel and transferred onto polyvinylidene fluoride for fluorescence membranes. Odyssey blocking buffer (LICOR Biosciences, Lincoln, NE, USA) was then added and membranes were blocked for 90min. Membranes were then probed with a mixture of a mouse anti-phospho-ERK1/2 antibody (1:2500; Sigma) and rabbit anti-ERK1/2 antibody (1:40000; Sigma) for 2–3h. Bands were visualized by the addition of a mixture of IRDye 800 (anti-mouse) antibody (1:10000; Sigma) and IRDye 680 (anti-rabbit) antibody (1:10000; Sigma) for 1h and scanned by the Odyssey infrared scanner (LICOR Biosciences). Bands densities were quantified using the scanner software and exported to Excel (Microsoft, Redmond, WA, USA). The level of phosphorylated ERK1/2 isoforms was normalized for differences in loading using the total ERK protein band intensities.

In vivo microdialysis in rat striatum

Male Sprague-Dawley rats (Charles River Laboratory, Wilmington, MA, USA), weighing 300–350g were used. Concentric microdialysis probes with 2mm long dialysis membranes were prepared as described previously.15 Animals were anesthetized with Equithesin (NIDA Pharmacy, Baltimore, MD, USA) and microdialysis probes were implanted in the ventral striatum (core of the nucleus accumbens); coordinates with respect to bregma: A 1.7, L +1.2 and V −7.6mm. The experiments were performed on freely moving rats 24h after the probe implantation. A Ringer solution (in mmoll−1) of 147 NaCl, 4 KCl and 2.2 CaCl2 was pumped through the dialysis probe at a constant rate of 1μl per minute. After a washout period of 90min, samples were collected at 20min intervals and split into two fractions of 10μl, to separately measure glutamate and dopamine contents. Each animal was used to study the effect of one treatment by local administration (perfusion by reverse dialysis) of the D4 receptor agonist RO-10-5824 or the D4 receptor antagonist L-745870. At the end of the experiment, rats were killed with an overdose of Equithesin and methylene blue was perfused through the probe. The brain was removed and placed in a 10% formaldehyde solution, and coronal sections were cut to verify the probe location. Dopamine content was measured by reverse high-performance liquid chromatography coupled to an electrochemical detector, as described in detail previously. Glutamate content was measured by high-performance liquid chromatography coupled to a flourimetric detector, as described before.16 The limit of detection (which represents three times baseline noise levels) for dopamine and glutamate was 0.5 and 50nM, respectively. Dopamine and glutamate values were transformed as percentage of the mean of the three values before the stimulation and transformed values were statistically analyzed with one-way repeated measures analysis of variance followed by Newman–Keuls tests, to compare glutamate and dopamine values of the samples obtained after drug perfusion with those obtained just before drug perfusion.

Neurotransmitter release in rat striatal slices

Rat brain slices were obtained from male Wistar rats weighing 180–220g. After rapid killing of the rat, the brain was immersed in oxygenated ice-cold artificial cerebrospinal fluid (ACSF) solution, and coronal brain slices (300μm thick) were obtained with a vibratome. The striatum (caudate-putamen and nucleus accumbens) was microdissected under a stereoscopic microscope and the slices were incubated for 30min at 37°C in ACSF (in mM: NaCl 118.25, KCl 1.75, MgSO4 1, KH2PO4 1.25, NaHCO3 25, CaCl2 2 and D-glucose 10), gassed continuously with O2/CO2 (95:5, v/v). For γ-aminobutyric acid (GABA) release, the slices were then incubated for 30min with 8nM [3H]GABA in 2ml solution containing 10μM aminooxyacetic acid (to inhibit GABA transaminase, thus preventing degradation of the labeled GABA). At the end of this period, excess radiolabeled compound was removed by washing twice with ACSF containing, in addition to aminooxyacetic acid and 10μM nipecotic acid (to prevent the reuptake of the released [3H]GABA). Both compounds were present in the perfusion solution for the rest of the experiment. For dopamine release, the slices were labeled with 77nM [3H]dopamine in Krebs–Henseleit solution containing 10μM pargyline, 0.57mM ascorbic acid and 0.03mM EDTA, which were present in the solutions for the rest of the experiment. For glutamate release, the tissues were incubated for 30min with 100nM [3H]glutamate in 2ml of ACSF containing 200μM aminooxyacetic acid (to inhibit glutamate decarboxylase and prevent the conversion of glutamate to GABA) and 200μM dihydrokainic acid (to prevent the uptake of [3H]glutamate by astrocytes). Dihydrokainic acid was present in the medium only during the incubation period. At the end of this period, the excess radiolabeled compound was removed by washing twice with ACSF. Methods for measuring [3H]neurotransmitter release and data analysis used in the present work were the same as those described previously.17, 18 The slices were apportioned randomly between the chambers (usually three slices per chamber) of a superfusion system (volume of each chamber 80μl; 20 chambers in parallel) and perfused with the ACSF at a flow rate of 0.5ml per minute for 1h. Basal release of [3H]neurotransmitter was measured by collecting four fractions of the superfusate (total volume 2ml) before depolarizing the slices with a solution in which the [K+] was raised to 25mM. The composition of the high K+ solution was (in mM): NaCl 101.25, KCl 23.75, MgSO4 1, KH2PO4 1.25, NaHCO3 25, CaCl2 2 and D-glucose 10. Six more fractions were collected in the high K+ medium. All drugs were added to the medium at fraction 2, before changing the superfusion to the high K+ medium, to explore effects on basal release. To determine the total amount of tritium remaining in the tissue, the slices were collected, treated with 1ml of 1M HCl and allowed to stand for 1h before adding the scintillator. The [3H]neurotransmitter release was expressed initially as a fraction of the total amount of tritium remaining in the tissue. The effect of drugs on the basal release of [3H]neurotransmitter was assessed by comparing the fractional release in fraction 2 (immediately before exposure of the tissue to the drug) and fraction four (immediately before exposure to 25mM of K+), using Student's paired t-test. Changes in depolarization-induced [3H]GABA release by drugs and treatments were assessed by comparing the area under the appropriate release curves between the first and last fractions collected after the change to high K+. The significance of drug effects was assessed by one-way analysis of variance and Tukey–Kramer test, using Prism Graph Pad Software 4.0 (Graph Pad Software). To obtain an unbiased estimate of IC50 values, concentration-response data were fitted by non-linear regression using the same software.

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Statistical analysis

Statistical analyses were performed with Prism Graph Pad Software 4.0 (Graph Pad Software). See above and figure legends (Figure 1 to Figure 7) for details.

Figure 1.
Figure 1 - Unfortunately we are unable to provide accessible alternative text for this. If you require assistance to access this image, please contact help@nature.com or the author

Human D2S and D4 receptors form heteromers in transfected cells. (a) Bioluminescence Resonance Energy Transfer (BRET) saturation curves were obtained from experiments with cells co-expressing, top to bottom, D2S-YFP (yellow fluorescent protein) and D4.2-RLuc (red), D4.4-RLuc (green) or D4.7-Rluc (blue), D2S-RLuc and D4.7-YFP (purple), A1-RLuc and D2S-YFP (black) or D4.4-RLuc and D1-YFP (gray). Co-transfections were performed with a constant amount of cDNA corresponding to the receptor-RLuc construct (2μg of cDNA for D4-RLuc or 1μg of cDNA for A1-RLuc) and increasing amounts of cDNA corresponding to the receptor-YFP construct (0.2–6μg of cDNA for D2S-YFP or 1–4μg of cDNA for D1-YFP). Both fluorescence and luminescence of each sample were measured before every experiment to confirm equal expression of Rluc (about 100000 luminescence units) while monitoring the increase of YFP expression (2000–20000 fluorescence units). BRET data are expressed as mean values±s.d. of four to nine different experiments grouped as a function of the amount of BRET acceptor. (b) BRET displacement experiments were performed in cells expressing constant amounts of D4.4-RLuc (2μg cDNA transfected) and D2S-YFP (2μg cDNA transfected) and increasing amounts (1–5μg of cDNA transfected) of D4.7 (blue) or D4.2 (green). Both fluorescence and luminescence of each sample were measured before every experiment to confirm no changes in the expression of D4.4-RLuc and D2S-YFP. BRET data are expressed as mean values±s.d. of five different experiments grouped as a function of the amount of BRET acceptor. Significant differences with respect to the samples without D4.2 or D4.7 were calculated by one-way analysis of variance (ANOVA) and Bonferroni's test (**P<0.01 and ***P<0.001). In (a, b), the relative amounts of BRET acceptor are expressed as the ratio between the fluorescence of the acceptor minus the fluorescence detected in cells only expressing the donor, and the luciferase activity of the donor. In the top, schematic representations of BRET (a) or BRET displacement (b) are shown. (c) Confocal microscopy images of cells transfected with 1μg of cDNA corresponding to, left to right, D4.2-RLuc, D4.4-RLuc or D4.7-RLuc and 0.5μg cDNA corresponding to D2S-YFP. Proteins were identified by fluorescence or by immunocytochemistry. D4-RLuc receptors are shown in red, D2S-YFP is shown in green and co-localization is shown in yellow. Scale bar: 5μm.

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Results

D2S and D4 receptors form heteromers in transfected cells

BRET experiments were performed where one of the receptor is fused to the bioluminescent protein Renilla Luciferase (RLuc) and the other receptor is fused to a YFP. The fusion proteins were functional (Supplementary Figure 1) and expressed at the membrane level (Figure 1c). Clear BRET saturation curves were obtained in cells expressing D4.2-RLuc or D4.4-RLuc receptors and increasing amounts of D2S-YFP (Figure 1a), but not in cells expressing D4.2-RLuc or D4.4-RLuc receptors and increasing amounts of D1-YFP (Figure 1a), indicating that the D4.2 and the D4.4 form heteromers with D2S but not with D1 receptors. Interestingly, in cells expressing the D4.7-Rluc variant and D1-YFP or D2S-YFP (Figure 1a) low linear BRET was detected, which was qualitatively similar to the results obtained with the negative control, with adenosine A1-RLuc and D2S-YFP receptors (Figure 1a). This result was not due to the particular BRET donor and acceptor chosen, as low and linear BRET were obtained when we swapped the fused proteins, that is, in cells co-expressing D2S-Rluc and D4.7-YFP (Figure 1a). These results strongly suggest that the human D4.7 polymorphic variant does not form heteromers with the human D2S receptor or if heteromers are formed, the fusion proteins are not properly oriented or are not within proximity to allow energy transfer (<10nm). One way to test if the receptors are indeed forming heteromers in such a way that impedes energy transfer is to titrate one receptor in the presence of the heteromer and look for changes in the BRET signal. In BRET displacement experiments, D4.2, but not D4.7 receptors were able to compete with D4.4-Rluc and alter heteromer formation with D2S-YFP (Figure 1b), meaning that D4.2 and D4.4, but not D4.7 receptors use the same molecular determinants to establish intermolecular interactions with D2S receptor and strongly suggesting that D4.7 receptors are unable to form heteromers with D2S.

D2S–D4 receptor heteromer signals through MAPK

To investigate the function of the D2S–D4 receptor heteromer, MAPK signaling (ERK1/2 phosphorylation) was determined. RO-10-5824 and quinelorane, selective D4 and D2/3 receptor agonists respectively,19, 20 selectively stimulated MAPK in cells transfected with D4 or D2S receptors, respectively (Supplementary Figure 2). Dose-response experiments with RO-10-5824 showed no significant differences between cells transfected with D4.2, D4.4 or D4.7 receptors (Supplementary Figure 2). However, in co-transfected cells, stimulation of D2S receptors potentiated D4 receptor-mediated MAPK activation, but not the other way around. Importantly, this functional interaction only occurred in cells transfected with D2S and D4.2 or D4.4, but not in cells expressing D4.7 receptors (Figure 2). Since disruption of D2S–D4 receptor heteromers (by substituting D4.2 or D4.4 with the D4.7 variant) is associated with the loss of the D2S–D4 receptor interaction at the MAPK level, this interaction constitutes a specific biochemical property of the D2S–D4 receptor heteromer and can be used as a biochemical fingerprint to detect the heteromer in native tissues.10

Figure 2.
Figure 2 - Unfortunately we are unable to provide accessible alternative text for this. If you require assistance to access this image, please contact help@nature.com or the author

Crosstalk between human D4 and D2S receptors in ERK1/2 phosphorylation. Cells were transiently co-transfected with 2.5μg of cDNA corresponding to D2S and 2.5μg of cDNA corresponding to D4.2 (a, d), D4.4 (b, e) or D4.7 (c, f). In (ac), cells were treated for 10min with increasing concentrations of RO-10-5824 in the presence () or in the absence (•) of quinelorane (50nM). In (df), cells were treated for 10min with increasing concentrations of quinelorane in the presence () or in the absence (•) of RO-10-5824 (50nM). The immunoreactive bands, corresponding to ERK1/2 phosphorylation, of three to six experiments were quantified and expressed as arbitrary units. For each curve, EC50 values were calculated as mean±s.e.m. and statistical differences between curves obtained in the presence or in the absence of quinelorane (ac) or RO-10-5824 (df) were determined by Student's t-test. EC50 with and without quinelorane: (a) 9±1 and 26±1nM (P<0.01), (b) 7±1 and 23±1nM (P<0.01), (c) 18±1 and 22±1nM (N.S.). EC50 with and without RO-10-5824: (d) 22±1 and 20±1nM (N.S.), (e) 20±1 and 17±1nM (N.S.), (f) 18±1 and 13±1nM (N.S.). N.S., non-statistical differences.

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D2S–D4 receptor heteromers in the mouse brain

D4 receptors are preferentially expressed in limbic areas and the prefrontal cortex, where they can be found in interneurons and also projecting neurons.1 In corticostriatal neurons, D4 receptors have also been localized at their nerve terminals,2, 3 where they can co-localize with D2S receptors.13 We therefore investigated the existence of D2S–D4 receptor heteromers in the striatum. Biophysical techniques cannot be easily applied in native tissues, but indirect methods can be used, such as the identification of a biochemical property of the heteromer (biochemical fingerprint).10 In this case, the biochemical fingerprint would be the potentiation by D2S receptor activation of D4 receptor-mediated MAPK activation, which should not occur with the human D4.7 variant. Before these experiments with mouse brain, we demonstrated by BRET saturation experiments in transfected cells that the mouse D2S receptor forms heteromers with the mouse D4 receptor (which has an amino-acid sequence in the 3IL similar to that from the human D4.2). Mouse fusion proteins were expressed in the plasma membrane of transfected cells (Figure 3a) and shown to be functional (Supplementary Figure 3). Like the human receptors, mouse D2S receptors were found to form heteromers with mouse D4 receptors and also with human D4.4 receptors, but not with human D4.7 receptors (Figure 3b). Furthermore, it was also shown that, in co-transfected cells, stimulation of the mouse D2S receptor potentiates the effect of the mouse D4, but not the human D4.7, on MAPK signaling (Figures 3c and d). This result was not reciprocal (Supplementary Figure 4) and mirrors the results obtained with human D4 and D2S receptors (Figure 2). We next analyzed the effects of D2 and D4 receptor agonists on MAPK signaling on striatal slices taken from knockin mice carrying the 7 repeats of the human D4.7 in replacement of the mouse region and from WT littermates (Figure 4). Neither quinelorane nor RO-10-5824 induced a significant ERK1/2 phosphorylation in striatal slices of WT mice when administered alone, but co-administration of both agonists produced a significant dose-dependent effect with an increase of up to fourfold (Figure 3e). This synergistic interaction between D2 and D4 receptors, which constitutes the biochemical fingerprint of the D2S–D4 receptor heteromer, was completely absent in the D4.7 mutant mouse (Figure 3e), confirming both the existence of D2S–D4 receptor heteromers and the absence of functional interactions between D2 and D4.7 receptors in the brain.

Figure 3.
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D2s–D4 receptor heteromers in the mouse brain. (a) Confocal microscopy images of cells transfected with 1μg of cDNA corresponding to, left to right, mouse D4-RLuc, human D4.4-RLuc and human D4.7-RLuc and 0.5μg of cDNA corresponding to D2S- yellow fluorescent protein (YFP). Proteins were identified by fluorescence or by immunocytochemistry. D4-RLuc receptors are shown in red, D2S-YFP is shown in green and co-localization is shown in yellow. Scale bar: 5μm. (b) Mouse D2S receptor heteromerization with mouse and human D4 receptors. Bioluminescence Resonance Energy Transfer (BRET) saturation curves were obtained from cells co-expressing mouse D4-Rluc (green), human D4.4-RLuc (red), human D4.7-RLuc (blue) or human A1-RLuc (gray) and mouse D2S-YFP receptors. Co-transfections were performed with a constant amount of cDNA corresponding to the receptor-RLuc construct (2μg of cDNA for mouse D4-RLuc, 2.5μg of cDNA for human D4-RLuc or 1μg of cDNA for A1-RLuc) and increasing amounts of cDNA corresponding to the receptor-YFP construct (0.2–6μg cDNA). Both fluorescence and luminescence of each sample were measured before every experiment to confirm equal expression of Rluc (about 100000 luminescence units) while monitoring the increase of YFP expression (2000–20000 fluorescence units). The relative amounts of BRET acceptor are expressed as the ratio between the fluorescence of the acceptor minus the fluorescence detected in cells only expressing the donor, and the luciferase activity of the donor. BRET data are expressed as mean values±s.d. of three to six different experiments grouped as a function of the amount of BRET acceptor. (c, d) Crosstalk between mouse D2S receptors and mouse or human D4 receptors in ERK1/2 phosphorylation. Cells transiently co-expressing mouse D2S receptors and mouse D4 receptors (c) or human D4.7 receptors (d) were treated for 10min with increasing RO-10-5824 concentrations in the presence () or in the absence (•) of quinelorane (50nM) before the ERK1/2 phosphorylation determination. The immunoreactive bands of three experiments (mean±s.e.m.; n=3) were quantified and expressed as arbitrary units. EC50 values with or without quinelorane were: (c) 7±0.1 and 15±0.1nM (Student's t-test: P<0.01) or (d) 18±0.1 and 15±0.1nM (Student's t-test: N.S.). (e) Striatal slices from wild-type (WT) or D4.7 mutant mice were treated for 10min with the indicated concentrations of RO-10-5824 (orange) or quinelorane (green) or with RO-10-5824 plus quinelorane (blue) and ERK1/2 phosphorylation was determined. For each treatment, the immunoreactive bands from four to six slices from a total 10 WT and 10 D4.7 mutant animals were quantified and values represent the mean±s.e.m. of the percentage of phosphorylation relative to basal levels found in untreated slices (100%). No significant differences were obtained between the basal levels of the WT and the D4.7 mutant mice. Significant treatment and genotype effects were shown by a bifactorial analysis of variance (ANOVA) followed by post hoc Bonferroni's tests (**P<0.01 and ***P<0.001, as compared with the lowest concentration of RO-10-5824).

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Figure 4.
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Targeted insertion of human variable number of tandem repeats (VNTRs) carrying 7 repeats into the mouse Drd4 exon 3 by homologous recombination in ES cells. (a) Structure of the Drd4 locus, targeting vector and targeted allele. (b) Southern blot analysis detected double homologous recombination events at the 5′ and 3′ ends using external probes after digestion with BamHI or EcoRI. (c) The presence of inserted human VNTR was verified by PCR using mouse primers flanking the expansion.

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D2–D4 receptor interactions modulate striatal glutamate release

To investigate the functional significance of D4 receptor activation, we determined D4 receptor-mediated modulation of striatal glutamate release by in vivo microdialysis in freely moving rats. The local perfusion of the D4 receptor agonist RO-10-5824 in the ventral striatum (in the nucleus accumbens) produced a dose-dependent decrease in the striatal extracellular concentration of glutamate and a concomitant increase in the extracellular concentration of dopamine (Figures 5a and 5b), which were counteracted by co-perfusion with the selective D4 receptor antagonist L-745870 (which was inactive when perfused alone) (Figures 5a–c). These results suggest that inhibitory D4 receptors are located in glutamatergic terminals, whose activation decreases basal striatal glutamate release. The increase in dopamine concentration can best be explained by a decreased activation of striatal GABAergic efferent neurons that tonically inhibit dopaminergic mesencephalic neurons. This interpretation could be confirmed in experiments with striatal slices, where dopamine should not be modified due to the interruption of the striatal-mesencephalic loop. In fact, in slices of dorsal or ventral rat striatum, the D4 receptor agonist RO-10-5824 decreased K+-induced glutamate release, an effect that was counteracted by the selective D4 receptor antagonist L-745870, but did not change dopamine or GABA release (Figure 6), indicating that striatal D4 receptors selectively and locally modulate glutamate release. This role of D4 receptors in the striatum can also explain previous results obtained with D4 receptor KO mice, which show an increase and decrease in the striatal extracellular concentration of glutamate and dopamine, respectively.21, 22

Figure 5.
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In vivo D4 receptor-mediated modulation of basal extracellular levels of glutamate in the rat ventral striatum. Effects of the local perfusion with the D4 receptor agonist RO-10-5824 and the D4 receptor antagonist L-745870 on the basal extracellular concentrations of glutamate (GLU) and dopamine (DA) in the ventral striatum (core of the nucleus accumbens). Horizontal bars show the periods of drug perfusion (concentrations are indicated in M). Data represent mean values±s.e.m. of the percentage of the mean of the three basal values before the first drug perfusion (n=6–8 per group): *P<0.05 and **P<0.01, compared with the values previous in time ‘0’ (repeated measures analysis of variance (ANOVA) followed by Newman–Keuls tests).

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Figure 6.
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D4 receptor-mediated modulation of [3H]glutamate, but not [3H]dopamine or [3H]GABA release from slices of dorsal and ventral striatum. Slices from the dorsal striatum (caudate-putamen; a, c, e) or the ventral striatum (nucleus accumbens; b, d, f) of reserpine-treated rats were treated with the D4 receptor agonist RO-10-5824 (100nM) or with the D4 receptor antagonist L-745870 (10nM) alone or in combination and the time course of K+-stimulated [3H]glutamate (a, b), [3H]dopamine (c, d) or [3H]GABA (e, f) release was determined. The RO-10-5824-induced effect (open circles) was prevented by the antagonist L-745870 (dark squares), which itself had no effect (open squares). Values are mean±s.e.m. of samples from three different animals performed in four replicates. Drug effect was assessed by comparing the relative area under the curve for each condition. **P<0.01 with respect to the control (analysis of variance (ANOVA) followed by Tukey–Kramer multiple comparison post hoc test).

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As mentioned before, there is evidence for co-localization of both D2 and D4 receptors in corticostriatal glutamatergic terminals2, 3, 13 and previous studies have demonstrated that presynaptic D2-like receptors have an inhibitory role in the modulation of striatal glutamate release.13, 23 However, since those studies did not use selective compounds, they could not distinguish between effects due to D2 or D4 receptor stimulation. Therefore, in this study we tested the effect of quinelorane alone and in combination with RO-10-5824 on glutamate release in rat striatal slices. To eliminate endogenous dopamine, rats were treated with reserpine, and the experiments performed in the presence of the D1-like receptor antagonist SCH-23390. Quinelorane significantly decreased K+-induced glutamate release, whereas the co-application of quinelorane with RO-10-5824 showed a more significant effect (Figure 7a). Dopamine strongly decreased K+-induced glutamate release, an effect partially counteracted by the D2 receptor antagonist L-741626 or by the D4 receptor antagonist L-745870, but completely counteracted by the simultaneous application of both antagonists (Figure 7b). In agreement with the reported higher in vitro affinity of D4 versus D2 receptor for dopamine,24 the IC50 of dopamine-mediated inhibition of K+-induced glutamate release was significantly higher in the presence of the D4 receptor antagonist (D2-mediated effect) than in the presence of the D2 receptor antagonist (D4-mediated effect) (Figure 7b). Finally, and more importantly, the D2 receptor agonist quinelorane synergistically potentiated the inhibitory effect of the D4 receptor agonist RO-10-5824 on K+-induced glutamate release (significant decrease in IC50 value) (Figure 7c), but not the other way around (Figure 7d). These results therefore show the same kind of D2–D4 receptor interaction demonstrated by D2S–D4 receptor heteromers in transfected cells with MAPK signaling. Our combined in vitro and in vivo data strongly suggest that D2S–D4 receptor heteromers are likely to have a key role in dopamine-mediated modulation of striatal glutamate release.

Figure 7.
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D2 and D4 receptor interactions in the modulation of striatal [3H]glutamate release. Striatal slices (dorsal striatum) from reserpine-treated rats were incubated with SCH-23390 (100nM) to block D1 receptor activation. In (a), slices were treated for 32min (fraction 2 to fraction 10) with medium (control), with the D4 receptor agonist RO-10-5824 (100nM), with the D2/3 receptor agonist quinelorane (100nM) or with both and K+-stimulated [3H]glutamate release was determined. Values are mean±s.e.m. of samples from three different animals performed in four replicates. Drug effects were assessed by comparing the relative area under the curve for each condition. **P<0.01 and ***P<0.001 with respect to the control and ##P<0.01 with respect to slices treated with RO-10-5824 or quinelorane alone (analysis of variance (ANOVA) followed by Tukey–Kramer multiple comparison post hoc test). In (b), slices were treated for 32min with increasing dopamine concentrations in the absence (dark circles) or in the presence of the D4 receptor antagonist L-45870 (10nM, dark squares), the D2 receptor antagonist L-741626 (10nM, open circles) or both (open squares) and K+-stimulated [3H]glutamate release was determined. Values are mean±s.e.m. of samples from three different animals performed in four replicates. Drug effects were assessed by comparing the relative area under the curve for each condition. The IC50 values were: 25.25nM (C.I.: 9.63–66.20nM) for dopamine alone, 5.75nM (2.12–15nM) for dopamine in the presence of L-741626 and 357.27nM (C.I.: 73.40–1739nM) for dopamine in the presence of L-745870. In (c), slices were treated for 32min with increasing concentrations of RO-10-5824 in the absence (black circles) or in the presence (open circles) of quinelorane (10nM) and K+-stimulated [3H]glutamate release was determined. In (d), slices were treated for 32min with increasing concentrations of quinelorane in the absence (black circles) or in the presence (open circles) of RO-10-5824 (10nM) and K+-stimulated [3H]glutamate release was determined. In (c, d), values are mean±s.e.m. of samples from three different animals performed in four replicates. The IC50 values were (c) 15nM (35.15–6.55nM) for RO-10-5824 alone and 0.05nM (1.21–0.02nM) for RO-10-5824 in the presence of quinelorane (Student's t-test: P<0.01) and (d) 2.55nM (7.31–0.89nM) for quinelorane alone and 1.48nM (4.5–0.45nM) for quinelorane in the presence of RO-10-5824 (Student's t-test; N.S.).

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Discussion

The present study shows that dopamine D2S and D4.2 or D4.4 receptors, but not the ADHD-associated human D4.7 variant, form functional heteromers in transfected cells and in the rodent brain. Co-stimulation of D2S and D4 receptors in the D2S–D4 receptor heteromer has a synergistic effect on MAPK signaling, which could be demonstrated in transfected cells and in the mouse striatum, but not in cells expressing D4.7 or in the striatum of a mutant mouse carrying the 7 repeats of the human D4.7 in the 3IL of the D4 receptor. These results provide a significant functional difference of one of the human receptor variants, D4.7, compared with the D4.2 and D4.4 variants, which can have important implications for the understanding of the pathogenesis of ADHD. Importantly, we also demonstrated, for the first time, that D2S–D4 receptor interactions modulate striatal glutamate release, suggesting that the D2S–D4 receptor heteromer allows dopamine to fine-tune glutamate neurotransmission.

The molecular mechanism involved in preventing heteromer formation between D2S and D4.7 receptors is not yet known. Indeed, the control of heteromer formation between G-protein-coupled receptors is still a large question in the field. Since the D4.7 receptor variant has the longest 3IL and is the only polymorphic form not forming heteromers with the D2S receptor, steric hindrance of the 3IL of D4.7 receptor is a probable mechanism responsible for this lack of heteromerization, but other mechanisms cannot be ruled out. Using two-hybrid methodologies as well as proteomic studies, interactions between dopamine receptors and a cohort of DRIPs (dopamine receptor interacting proteins) have been demonstrated, forming signaling complexes or signalplexes.25, 26 Some of these DRIPs show selectivity for some dopamine receptor subtypes. For example, filamin or protein 4.1N interact with D2 and D3 receptors but not with D1, D5 or D4 receptors,27, 28 the PDZ domain-containing protein, GIPC (GAIP interacting protein, C terminus) interacts with D2 and D3 receptor but not with the D4 receptor subtype29 and paralemmin interacts exclusively with D3, but not with D2 or D4 receptors.30 All of these interactions modulate receptor targeting, trafficking and signaling. Proline-rich sequences of the D4 receptor, mainly located in the polymorphic region of the 3IL, constitute putative SH3 binding domains, which can potentially interact with adapter proteins like Grb2 and Nck, which do not have any known catalytic activity but are capable of recruiting multiprotein complexes to the receptor.24 It can be hypothetized that differences in DRIPs recruitment by D4.7 and the other D4 polymorphic forms can influence the D4.7 ability to form heteromers, but future studies will be required.

Previous experiments indicated that locally in the striatum, dopamine inhibits glutamate release by activating D2 receptors (predominantly D2S) localized in glutamatergic terminals.13, 15 Other studies also indicate that striatal postsynaptic D2 receptors (predominantly D2L) indirectly modulate glutamate release by retrograde endocannabinoids signaling.31 The present results indicate that D4 receptors also have a key role in the modulation of striatal glutamate release, likely through its ability to form heteromers with presynaptic D2S receptors. In the striatal D2S–D4 receptor heteromer, low concentrations of dopamine should bind to the D4 receptor, which has more affinity for dopamine than the D2S receptor,24 causing a certain degree of inhibition of glutamate release. However, at higher concentrations, dopamine should also bind to the D2S receptor and under these conditions, the synergistic interaction in the D2S–D4 receptor heteromer will produce an even stronger inhibition of glutamate release. Therefore, the D2S–D4 receptor heteromer seems to act as a concentration-dependent device that establishes two different degrees of presynaptic dopaminergic control over striatal glutamatergic neurotransmission. Since the strong modulation observed with higher concentrations of dopamine depends on D2S–D4 receptor heteromerization, the existence of a D4.7 variant implies a weaker control of glutamatergic neurotransmission, which could be a main mechanism involved in the pathogenesis of ADHD. This could also explain at least part of the so far not understood successful effect of psychostimulants in ADHD, which amplify dopaminergic signaling and these medications appear to be more effective in ADHD patients with the D4.4 than with the D4.7 variants.32, 33 We have to take into account that the existence of a D4.7 variant does not imply ADHD is the result of this variant, but rather that it is one factor that contributes to its development. In fact, the D4.7 variant might constitute a successful evolutionary trait under the appropriate environmental exposure.7, 34 The present study provides a new element of interest in the field of receptor heteromes, which now become new targets to be studied when dealing with functional differences associated with polymorphisms of G-protein-coupled receptor genes.

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Conflict of interest

The authors declare no conflict of interest.

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Acknowledgements

We thank the technical help from Jasmina Jiménez (University of Barcelona). The study was supported by the NIDA IRP funds and from Grants from Spanish Ministerio de Ciencia y Tecnología (SAF2008-03229-E, SAF2009-07276, SAF2010-18472, SAF2008-01462 and Consolider-Ingenio CSD2008-00005) and from Consejo Nacional de Ciencia y Tecnología de México (50428-M). PJM is a Ramón y Cajal Fellow.

Supplementary Information accompanies the paper on the Molecular Psychiatry website