Cannabinoid CB1 Receptor Activation Mitigates N‑Methyl‑d‑aspartate Receptor-Mediated Neurotoxicity
Centro de Investigación Biomédica en Red Enfermedades Neurodegenerativas (CiberNed), National Institute of Health Carlos III, 28031 Madrid, Spain
Institut de Neurociències UB, Campus Mundet, 08035 Barcelona, Spain
Department of Biochemistry and Physiology, Faculty of Pharmacy and Food Science, 16724University of Barcelona, 08028 Barcelona, Spain
Departament de Biochemistry and Molecular Biomedicine, 16724University of Barcelona, 08028 Barcelona, Spain
Institut de Neurociències, Department de Bioquímica i Biologia Molecular, Facultat de Medicina, Universitat Autònoma de Barcelona, Bellaterra, 08193 Barcelona, Spain
Departament de Bioquímica i de Biologia Molecular, Universitat Autònoma de Barcelona, Bellaterra, 08193 Barcelona, Spain
Centro Singular de Investigación en Química Biolóxica e Materiais Moleculares (CiQUS), 15705 Santiago de Compostela, Spain
Institut de Química Teòrica i Computacional (IQTCUB), School of Chemistry, University of Barcelona, 08028 Barcelona, Spain
*Email: g.navarro@ub.edu. *Email: rfranco@ub.edu.Abstract
Alzheimer’s disease (AD) is characterized by synaptic dysfunction and excitotoxicity, yet effective therapeutic strategies remain limited. This study explores the functional and physical interplay between cannabinoid CB1 receptors (CB1Rs) and N-methyl-d-aspartate receptors (NMDARs), which are implicated in AD pathology. Using bioluminescence resonance energy transfer and imaging assays in HEK-293T cells, we demonstrate a direct interaction between CB1R and the N1 subunit of NMDAR, supporting the formation of receptor complexes. Functional assays further reveal a bidirectional negative crosstalk: NMDA attenuates CB1R-mediated cAMP inhibition, while CB1R activation reduces NMDA-induced calcium influx and mitogen-activated protein kinase signaling pathway activation. This negative crosstalk suggests the existence of receptor–receptor interactions with functional consequences. Complexes of CB1Rs and N1 subunits of NMDARs are present in both neurons and microglia, and their expression is upregulated in response to Aβ1–42 and in cells derived from the APPSw/Ind AD model mice. However, upregulation did not always correlate with stronger CB1R–NMDAR cross-modulation, suggesting that cell-specific signalosome composition shapes the signaling outcome. Functionally, CB1R activation confers neuroprotection: It rescues neurite loss induced by NMDA and Aβ1–42, highlighting the therapeutic potential of modulating CB1R–NMDAR interactions. These findings support a model in which CB1R–NMDAR interactions, through dynamic functional cross-modulation, finely tune excitotoxic and inflammatory signaling pathways. This mechanism offers therapeutic prospects for addressing cannabinoid-glutamatergic interactions.
Alzheimer’s disease (AD) was first described by Alois Alzheimer in 1906 and nowadays represents 60 to 80% of the 55 million cases of dementia around the world.ref1 There are two forms of AD; the sporadic form is characterized by a not fully understood etiopathogenesis and by the influence of different factors such as lifestyle, environment, and other genetic and epigenetic alterations. It accounts for about 95% of AD cases. The familial AD is caused by mutations in the amyloid precursor protein (APP) and in the presenilin-1 and -2 genes and represents around 5% of cases.ref2 The fact that AD still does neither have a cure not interventions to delay disease progression leads to the necessity of discovering new therapeutic targets.
AD begins with the loss of ability to retain new memories and progresses with different cognitive and behavioral changes, which worsen over time. The pathophysiology of AD is characterized by aggregates of amyloid precursor protein (APP)-derived ß-amyloid (Aβ) peptides and hyperphosphorylated tau protein. ref3,ref4 Overall, synaptic dysfunction and neuronal death generally result in progressive neurodegeneration.ref5 Neurodegeneration is accompanied by alterations in synaptic plasticity that, at least, is partially mediated by altered glutamatergic neurotransmission. In fact, N-methyl-d-aspartate (NMDA) and α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA) glutamate receptors are directly involved in synaptic plasticity.ref6
The NMDA glutamate receptor (NMDAR) is an ion channel that requires the binding of two coagonists for activation: glutamate and either glycine or d-serine. The receptor is typically formed by a tetramer consisting of two N1 and two N2 subunits. Glutamate binds to the N2 subunits, while glycine or d-serine bind to the N1 subunits. In addition to ligand binding, channel opening also requires relief of a voltage-dependent magnesium block, which occurs upon membrane depolarization.ref7 Glutamate is the main excitatory neurotransmitter in the human brain. However, sustained elevations in extracellular glutamate levels can lead to excitotoxicity and neurodegeneration. Synaptic NMDARs, primarily containing N2A subunits, are generally associated with cell survival and neuroprotective signaling, whereas extrasynaptic NMDARs, often containing N2B subunits, are linked to neuronal vulnerability and excitotoxic pathways.ref8
NMDAR function may be regulated by neurotransmitters or neuromodulators acting via G protein-coupled receptors (GPCRS). ref9−ref10 ref11 The activation of adenosine A2A receptors modulates NMDAR functionality likely by a direct interaction of NMDAR and adenosine A2A receptors.ref10 NMDARs may also interact with dopamine D1 and histamine H3 receptors, forming NMDAR-D1R-H3R functional complexes.ref9 The regulation of NMDARs by interaction with cannabinoid receptors is relevant in the AD context. NMDAR may interact with the cannabinoid CB2 receptor (CB2R), and evidence of the interaction was found both in primary cultures of neurons and microglia and in brain sections.ref12 The print of NMDAR–CB2R interaction consists of an impairment of NMDAR function by CB2R activation. This feature was detected in microglial cells activated with LPS plus IFN-γ, and in primary microglia from the brain of APPSw/Ind mice, a transgenic model of AD.ref13
Cannabinoids bind to two different receptors: CB1 and CB2. In the central nervous system, CB2Rs are mainly expressed in glia, whereas CB1 receptors (CB1Rs) are expressed in both neurons and glia. In humans, the cannabinoid CB1R is primarily expressed in the cortex, hippocampus, cerebellum, basal ganglia, and brainstem.ref14 Although its pathophysiological role in AD remains unclear, the genetic deletion of the CB1Rs associates with a loss of neurons and a cognitive deficit in the hippocampus of adult KO mice.ref15 Interestingly, acute activation of CB1R, particularly at a young age, impairs short-term memory in a dose-dependent manner.ref16 Multiple studies have demonstrated that CB1Rs can counterbalance the effects of NMDA receptors (NMDARs), thereby contributing to the neuroprotective role of cannabinoids against excessive NMDAR activation. ref17−ref18 ref19 This interaction plays a role in modulating nociceptive signaling under glutamatergic stimulation, as disruption of CB1R function has been shown to result in heightened NMDAR-mediated pain responses, including allodynia and hyperalgesia.ref20 Importantly, in neuropathic conditions characterized by NMDAR overactivity, which typically reduces the efficacy of potent analgesics like opioids, cannabinoids may still retain partial analgesic activity.ref21
While CB1Rs are well-known for their presynaptic localization, they are also present postsynaptically,ref22 where they overlap with the expression of NMDARs. In fact, N1 subunits of NMDARs colocalize with CB1Rs on neuronal somata and dendrites in the rat hippocampus.ref23 In addition, Sánchez-Blázquez et al. reported that CB1R coprecipitates with the N1 subunit, but notor only minimallywith the N2 subunit. Notably, this interaction was reduced following NMDAR overactivation.ref24 Here, we demonstrate a molecular interaction between the CB1R and subunits of the NMDAR in transfected HEK-293T cells, primary neurons and primary microglia. Functional signaling data, such as ERK1/2 phosphorylation and calcium responses, obtained in these cells as well as in primary microglia and neurons, show a crosstalk between CB1 and NMDA receptors. Furthermore, the characteristics of the inter-receptor modulation vary from cell to cell and are context dependent.
In this study, we aimed to investigate the molecular and functional interaction between CB1 receptors and NMDARs in neurons and microglia, with a particular focus on their modulation by Aβ1‑42 and in the context of AD. By combining biochemical, imaging, and functional approaches, we sought to determine how CB1R–NMDAR complexes contribute to the regulation of excitotoxic and inflammatory signaling pathways, and to evaluate their potential as neuroprotective targets.
Materials and Methods
Reagents
LPS, interferon-γ (IFN-γ), and Aß1–42 peptide were purchased from Sigma-Aldrich (St Louis, Missouri), and ACEA, rimonabant, NMDA, and MK-801 were purchased from Tocris Bioscience (Bristol, UK).
Transgenic Mouse Model of Alzheimer’s Disease (AD)
APPSw/Ind transgenic mice (line J9; C57BL/6 background) expressing human APP695 harboring the familial Alzheimer′s disease-linked Swedish (K670N/M671L) and Indiana (V717F) mutations under the PDGFβ promoter were obtained by crossing heterozygous APPSw/Ind with nontransgenic animals.ref25 Mice at 2 days of age were genotyped individually by conventional PCR. ref26,ref27 Experimental procedures were conducted according to the Animal and Human Ethical Committee of the Universitat Autònoma de Barcelona (protocol CEEAH 1783, Generalitat Catalunya 6381) following the European Union guidelines. Experiments with primary cultures (see below) were performed blindly, without knowing the genotype, which was disclosed for data analysis.
Aß1–42 Oligomer Production
Human Aß-oligomers (Sigma-Aldrich, St Louis, Missouri) were prepared according to previously established protocol.ref28 Briefly, the lyophilized Aß1–42 was disaggregated in 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP, Sigma-Aldrich, St Louis, Missouri, USA, Cat. No. B2517) to 0.5 mg/mL. Then, the HFIP was evaporated at room temperature and stored at −80 °C. 24 h before the experiment, the peptide was dissolved in DMSO (Sigma-Aldrich, 1/1000) and then sonicated. Finally, the peptide was diluted to 50 μM in F12 medium (Gibco) and kept at 4 °C for 24 h to allow oligomerization. For each experiment, a fresh oligomer preparation was used.
Expression Vectors
The cDNA for the human version of cannabinoid CB1R without its stop codon was obtained by PCR and subcloned to a SNAP-containing vector (PSNAP; Cisbio Bioassays) and YFP-containing vector (pEYFP-N1, PerkinElmer, Wellesley, Massachusetts) using sense and antisense primers harboring unique restriction sites for HindIII and BamHI, generating the SNAP- and YFP-tagged CB1R (SNAP-CB1R and CB1R-YFP). The cDNA for the human version of the N1 subunit of the NMDA receptor without its stop codon was obtained by PCR and subcloned to the Rluc-containing vector (pRluc-N1; PerkinElmer, Wellesley, Massachusetts) using sense and antisense primers harboring unique restriction sites for HindIII and BamHI, generating N1-Rluc.
Cell Culture and Transfection
HEK-293T cells (lot #70022180) were acquired from the American Type Culture Collection (ATCC). Cells were amplified and grown until passage 18 in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2 mM l-glutamine, 100 U/mL penicillin/streptomycin, and 5% (v/v) heat-inactivated fetal bovine serum (FBS) (Invitrogen, Paisley, Scotland, UK). Cells were maintained in a humid atmosphere of 5% CO2 at 37 °C. Cells were transiently transfected with the PEI (polyethylenimine, Sigma, St. Louis, Missouri, USA) method, as previously described (Navarro et al., 2015). To prepare mouse primary microglia, the brain was removed from P2–P3 C57BL/6 mouse pups. Microglial cells were isolated as described in ref ref29 and plated at confluence of 40,000 cells/0.32 cm2 and grown in DMEM supplemented with 2 mM l-glutamine, 100 U/mL penicillin/streptomycin, and 5% (v/v) heat-inactivated fetal bovine serum (FBS) (Invitrogen, Paisley, Scotland, UK) for 12 days. For preparing primary neurons of mice, cortex and hippocampus were removed from the brain of from E18–E19 fetuses. Neurons were isolated as described in ref ref30 and plated at a confluence of 40,000 cells/0.32 cm2. Neuronal cells were grown in neurobasal medium supplemented with 2 mM l-glutamine, 100 U/mL penicillin/streptomycin, and 2% (v/v) B27 supplement (Gibco) in a 96-well plate for 12 days.
HTRF Assay
HTRF assays were performed in transfected HEK-293T cells expressing SNAP-tagged CB1R in the presence or in the absence of NMDAR. Tag-lite-based binding assays were performed 24 h after transfection. For cell labeling, culture medium was removed followed by the addition of 100 nM SNAP-Lumi4-Tb, previously diluted in 3 mL of TLB 1× for 1 h at 37 °C under a 5% CO2 atmosphere in a cell incubator. Cells were then washed four times with 2 mL of TLB 1× to remove the excess of SNAPLumi4-Tb, detached with enzyme-free cell dissociation buffer, centrifuged for 5 min at 1500 rpm and collected in 1 mL of TLB 1×. Densities in the 2500–3000 cells/well range were used to carry out binding assays in white opaque 384-well plates.
The fluorophore-conjugated CB1R ligand (labeled CELT-335) and unconjugated CB1R agonist, ACEA, were diluted in TLB 1×. HEK-293T cells transiently expressing Tb-labeled SNAP-CB1R with or without NMDAR were incubated with a 20 nM fluorophore-conjugated CB1R ligand, in the presence of increasing concentrations (0–10 mM range) of ACEA. Plates contained 10 mL of labeled cells, and 5 mL of TLB 1× or 5 mL of ACEA was added prior to the addition of 5 mL of the fluorescent ligand. Plates were then incubated for at least 2 h at room temperature before signal detection. A detailed description of the HTRF assay is found in ref ref31. Signal was detected using a PHERAstar FS (BMG Lab Technologies, Offenburg, Germany) microplate reader equipped with a FRET optic module allowing donor excitation at 337 nm and signal collection at both 665 and 620 nm. A frequency of 10 flashes/well was selected for the xenon flash lamp excitation. The signal was collected at both 665 and 620 nm using the following time-resolved settings: delay, 150 ms; integration time, 500 ms. HTRF ratios were obtained by dividing the acceptor (665 nm) by the donor (620 nm) signals and multiplying by 10,000. The 10,000-multiplying factor is used solely for the purpose of easier data handling.
Bioluminescence Resonance Energy Transfer (BRET) Assay
HEK-293T cells were transiently cotransfected with a constant amount of cDNA encoding for N1-Rluc and N2B in pcDNA3.1 or dopaminergic D1R-Rluc as negative control and with increasing amounts of cDNA corresponding to CB1R-YFP. 48 h after transfection, cells were adjusted to 20 μg of protein using a Bradford assay kit (Bio-Rad, Munich, Germany), using bovine serum albumin for standardization. To quantify protein-YFP expression, fluorescence was read in a Mithras LB 940 equipped with a high-energy xenon flash lamp, using a 30 nm bandwidth excitation filter at 485 nm reading. For BRET measurements, readings were collected 1 min after the addition of 5 μM coelenterazine H (Molecular Probes, Eugene, Oregon) using a Mithras LB 940, which allows the integration of the signals detected in the short-wavelength filter at 485 nm and the long-wavelength filter at 530 nm. To quantify protein-Rluc expression, luminescence readings were performed 10 min after 5 μM coelenterazine H addition using a Mithras LB 940.
Immunocytochemistry
HEK-293T cells were transfected with N1-Rluc, N2B, and CB1R-YFP, fixed in 4% paraformaldehyde for 15 min, and washed twice with PBS containing 20 mM glycine before permeabilization with PBS-glycine containing 0.2% Triton X-100 (5 min incubation). HEK-293T cells were treated for 1 h with PBS containing 1% bovine serum albumin, labeled with the primary mouse anti-Rluc antibody, and subsequently treated with a Cy3-conjugated antimouse (1/200; Jackson ImmunoResearch (red)) secondary antibody for 1 h. The CB1R-YFP fusion protein was detected by YFP’s own fluorescence. Colocalization is observed in yellow. Nuclei were labeled with Hoechst (blue). Samples were washed several times and mounted with 30% MOWIOL (Calbiochem). Samples were observed under a Zeiss 880 confocal microscope (Carl Zeiss, Oberkochen, Germany). Scale bar: 10 μm.
cAMP Level Determination
cAMP experiments were performed as previously described. ref32,ref33 HEK-293T cells were transfected with the cDNAs for N1 (0.5 μg/well), N2B (0.5 μg/well), and CB1R (0.3 μg/well). Two hours before initiating the experiment, HEK-293T, neuronal, or microglial cell culture medium was replaced by serum-starved DMEM medium. Then, cells were detached, resuspended in growing medium containing 50 mM zardaverine, and placed in 384-well microplates (2500 cells/well). Cells were pretreated (15 min) with selective antagonists (SR141617 for CB1R or MK-801 for NMDAR) followed by agonist stimulation (ACEA for CB1R and/or NMDA for NMDAR) or vehicle for an additional period of 15 min before adding 0.5 mM forskolin (FK) or vehicle. Readings were performed after 60 min of incubation at 25 °C. HTRF energy transfer measures were performed using the Lance Ultra cAMP kit (PerkinElmer, Waltham, Massachusetts, United States). Fluorescence at 665 nm was analyzed in a PHERAstar Flagship microplate reader equipped with an HTRF optical module (BMG Lab Technologies, Offenburg, Germany).
ERK Phosphorylation Assays
To determine the phosphorylation of extracellular signal-regulated kinase 1/2 (ERK1/2), 50,000 HEK-293T cells/well, neurons, or microglia were plated in transparent Deltalab 96-well microplates and kept at the incubator for 24 h. HEK-293T cells were transfected with the cDNAs for N1 (0.5 μg/well), N2B (0.5 μg/well), and CB1R (0.3 μg/well). 2 to 4 h before the experiment, the medium was substituted by serum-starved DMEM. Then, cells were pretreated at 25 °C for 10 min with selective antagonists (SR141617 for CB1R or MK-801 for NMDAR) followed by agonist stimulation (ACEA for CB1R and/or NMDA for NMDAR) or vehicle. Cells were then washed twice with cold PBS before the addition of lysis buffer (20 min treatment). Ten microliters of each supernatant was transferred to white ProxiPlate 384-well microplates, and ERK1/2 phosphorylation was quantified using the AlphaScreen SureFire Kit (PerkinElmer, Waltham, Massachusetts, USA), following the manufacturer’s instructions. Signal detection was performed using an EnSpire Multimode Plate Reader (PerkinElmer).
To assess ERK1/2 phosphorylation in hippocampal and cortical slices, tissues were maintained in HBSS supplemented with glucose for 2 h at 37 °C and then stimulated for 15 min with selective agonists. Samples were sonicated on ice (2 × 10 s pulses) in lysis buffer, and the total protein concentration was adjusted to 1 μg/μL using SDS and lysis buffer. Equal amounts of protein (20 μg) were separated by electrophoresis on 10% SDS–polyacrylamide gels and transferred to PVDF membranes (Immobilon-FL, Merck, St. Louis, Missouri, USA) using the Trans-Blot Turbo system (Bio-Rad) for 30 min. Membranes were blocked for 2 h at room temperature with constant shaking using Odyssey Blocking Buffer (LI-COR Biosciences, Lincoln, Nebraska, USA) and incubated overnight at 4 °C with a primary antibody mix containing mouse antiphospho-ERK1/2 (1:2500; Merck, ref. M8159) and rabbit antitotal ERK1/2 (1:40,000; Merck, ref. M5670). After washing with 0.05% PBS-Tween, membranes were incubated for 2 h at room temperature (protected from light) with IRDye 800CW antimouse (1:10,000; Merck, ref. 926-32210) and IRDye 680RD antirabbit (1:10,000; Merck, ref. 926-68071) secondary antibodies. Membranes were washed again with PBS-Tween, and bands were visualized using an Odyssey Infrared Imaging System (LI-COR Biosciences). Band intensities were quantified using Fiji software. Phospho-ERK1/2 levels were normalized to total ERK1/2, and results are expressed as percentage relative to basal (vehicle-treated) conditions.
Dynamic Mass Redistribution (DMR) Assays
Cell mass redistribution induced upon receptor activation was detected by illuminating the underside of a biosensor with polychromatic light and measuring the changes in the wavelength of the reflected monochromatic light. The magnitude of this wavelength shift (in picometers) is directly proportional to the amount of DMR. HEK-293T cells expressing the cDNAs for N1, N2B and CB1R were seeded in 384-well sensor microplates to obtain 70–80% confluent monolayers constituted by approximately 10,000 cells per well. Previous to the assay, cells were washed twice with assay buffer (HBSS with 20 mM HEPES, pH 7.15) and incubated for 2 h with assay buffer containing 0.1% DMSO (24 °C, 30 mL/well). Hereafter, the sensor plate was scanned and a baseline optical signature was recorded for 10 min before adding 10 mL of the selective antagonists (SR141617 for CB1R or MK-801 for NMDAR) for 30 min followed by the addition of 10 mL of specific agonists (ACEA for CB1R and/or NMDA for NMDAR) or vehicle; all test compounds were dissolved in assay buffer. The cell signaling signature was determined using an EnSpire Multimode Plate Reader (PerkinElmer, Waltham, Massachusetts, United States) by a label-free technology. Then, DMR responses were monitored for at least 5,000 s. Results were analyzed using an EnSpire Workstation Software v 4.10.
ß-Arrestin-2 Recruitment
ß-Arrestin-2 recruitment was determined as previously described.ref33 Briefly, BRET experiments were performed in HEK-293T cells 48 h after transfection with the cDNAs corresponding to the CB1R-YFP (1 μg cDNA), N1 (05 μg cDNA), N2B (0,5 μg cDNA), and ß-arrestin-2-Rluc (0,5 μg cDNA). Cells (20 mg protein) were distributed in 96-well microplates (Corning 3600, white plates with white bottom) and were incubated with 10 mL of selective antagonists (SR141617 for CB1R or MK-801 for NMDAR) for 10 min followed by the addition of 10 mL of specific agonists (ACEA for CB1R and/or NMDA for NMDAR) or vehicle 1 min before adding coelenterazine H (0.5 μM). BRET between ß-arrestin-2-Rluc and CB1R-YFP was immediately determined and quantified. The readings were collected using a Mithras LB 940 (Berthold Technologies, Bad Wildbad, Germany) that allows the integration of the signals detected in the short-wavelength filter at 485 nm and the long-wavelength filter at 530 nm. To quantify CB1R-Rluc expression, luminescence readings were also performed 10 min after adding 5 μM coelenterazine H.
Cytosolic Calcium Determination
HEK-293T cells were cotransfected with the cDNA for CB1R (0.3 μg/well), N1 (0.5 μg/well), N2B (0.5 μg/well), and 0.5 μg/well of the GCaMP6 calcium sensorref34 using PEI protocol (see Cell Culture and Transfection). 48 h after transfection, cells (150,000 HEK-293T cells/well in 96-well black, clear bottom microtiter plates) were incubated with Mg2+-free Locke’s buffer pH 7.4 (154 mM NaCl, 5.6 mM KCl, 3.6 mM NaHCO3, 2.3 mM CaCl2, 5.6 mM glucose, and 5 mM HEPES) supplemented with 10 μM glycine and receptor antagonists (SR141617 for CB1R or MK-801 for NMDAR) for 10 min followed by agonists stimulation (ACEA for CB1R and/or NMDA for NMDAR) just a few seconds before readings. The fluorescence emission intensity of GCaMP6 was recorded at 515 nm upon excitation at 488 nm on the EnSpire Multimode Plate Reader for 450 s every 15 s and 100 flashes per well.
Proximity Ligation Assays (PLA)
The interaction between NMDA and CB1R was detected using the Duolink II In Situ PLA Detection Kit (OLink; Bioscience, Uppsala, Sweden) following the instructions of the supplier. Primary neurons and microglial cells were grown on glass coverslips and were fixed in 4% paraformaldehyde for 15 min, washed with PBS containing 20 mM glycine to quench the aldehyde groups, permeabilized with the same buffer containing 0.05% Triton X-100 for 5 min, and successively washed with PBS. After 1 h of incubation at 37 °C with the blocking solution in a preheated humidity chamber, primary cells were incubated overnight in the antibody diluent medium with a mixture of equal amounts of rat monoclonal anti-N1 antibody (1:200, Millipore) and a polyclonal rabbit anti-CB1R antibody (1:100, Abcam). Cells were processed using the PLA probes detecting primary antibodies (Duolink II PLA probe plus and Duolink II PLA probe minus) diluted in the antibody diluent to a concentration of 1:5. Ligation and amplification were done as indicated by the supplier and cells were mounted using the mounting medium with Hoechst (1/200; Sigma). Samples were observed under a Zeiss 880 confocal microscope (Carl Zeiss, Oberkochen, Germany) equipped with an apochromatic 63× oil-immersion objective (N.A. 1.4) and 405 nm and 561 nm laser lines. For each field of view, a stack of two channels (one per staining) and four to eight Z stacks with a step size of 1 μm were acquired. A quantification of cells containing one or more red spots versus total cells (blue nucleus) and, in cells containing spots, the ratio r (number of red spots/cell) were determined. One-way ANOVA followed by Bonferroni’s post hoc multiple comparison test was used to compare the values (% of positive cells or r spots/cell) obtained for each pair of receptors.
Neurite Patterning Determination
Cortical and hippocampal primary neurons seeded for 11 days were treated with Aβ (500 nM) for 48 h and subsequently stimulated with NMDA (15 μM), ACEA (100 nM), or vehicle for 24 h. Then, cells were fixed in 4% paraformaldehyde for 15 min and then washed twice with PBS containing 20 mM glycine followed by permeabilization with the same buffer containing 0.2% Triton X-100 (15 min incubation). Samples were treated for 1 h with blocking solution (PBS containing 1% bovine serum albumin) and labeled with polyclonal rabbit anti-Nectin 3 antibody (Abcam, 1/1000). Neurons were detected with 3 anti-F-actin antibody fused to an Alexa 488 fluorophore (Thermo Fisher, 1/400). Then, samples were incubated at RT for 2 h with a Cy3-conjugated antirabbit secondary antibody (1/200, 711-166-152, Jackson ImmunoResearch). Finally, cells were washed several times with PBS and mounted with 30% MOWIOL (Calbiochem, San Diego, California, USA). Nuclei were stained with Hoechst (1/100). Samples were observed under a Zeiss 880 confocal microscope (Leica Microsystems, Wetzlar, Germany). Quantification of neurite formation was performed over segments of 15 μm. Each red dot represents a neurite formation.
Data Handling and Statistical Analysis
Data from homogeneous binding assays were analyzed using Prism 6 (GraphPad Software, Inc., San Diego, California, United States). K i values were determined according to the Cheng and Prusoff equation (Cheng, 2001). Signal-to- background (S/B ratio) calculations were performed by dividing the mean of the maximum value (μmax) by that of the minimum value (μmin) obtained from the sigmoid fits. The data are shown as the mean ± SEM. Statistical analysis was performed with Prism 8.0 software. Significance was analyzed by one-way ANOVA, followed by Bonferroni’s multiple comparison post hoc test. Significant differences were considered when p < 0.05.
Results
Cannabinoid CB1 Receptor Shows Lower Affinity for Its Agonist when Forming Complexes with NMDAR
Colocalization (in yellow) between CB1R and the N1 subunit of the NMDA receptor was detected in HEK-293T cells expressing CB1R-YFP, N1-Rluc, and N2B (Figure fig1 A). CB1R-YFP was detected by the YFP’s own fluorescence (shown in green), and the Cy3-conjugated antimouse secondary antibody was used to label the anti-Rluc primary antibody (shown in red). Next, Bioluminescence Resonance Energy Transfer (BRET) assays were performed in HEK-293T cells expressing a constant amount of N1-Rluc and N2B and increasing amounts of CB1R-YFP. A saturation BRET curve that demonstrates the physical interaction between the two proteins was obtained (Figure fig1 B). A similar experiment was performed in cells expressing a constant amount of dopamine D1 receptors fused to Rluc and increasing amounts of CB1R-YFP. The linear relationship observed (Figure fig1 B) indicates no interaction between the two proteins, validating the condition as a negative control.
One potential consequence of receptor–receptor interactions is an alteration in ligand affinity. To assess whether coexpression of the N1 and N2 subunits of the NMDAR affects the affinity of CB1R for its ligands, we performed a nonradioactive Homogeneous Time-Resolved FRET (HTRF) assay using CELT-335, a potent and selective fluorescent CB1R ligand.ref35 SNAP-tag technology (see Materials and Methods) was applied in HEK-293T cells expressing the SNAP protein fused to the CB1R. Under these conditions, the K i value of ACEA, a selective CB1R agonist competing with CELT-335 for binding, was 4.8 nMconsistent with previously reported values (Figure fig1 C). However, when the same assay was conducted in cells coexpressing SNAP-CB1R along with the N1 and N2 subunits of the NMDAR, the K i value for ACEA increased to 17.7 nM (Figure fig1 D). This reduced affinity suggests that the presence of NMDAR subunits induces conformational changes in CB1R, possibly due to direct receptor–receptor interaction. As demonstrated below, the coexpression of N1 and N2 subunits leads to functional outputs that support the notion that conformational changes in CB1Rs result from its interaction with fully assembled NMDA receptors.
Further Evidence of Receptor–Receptor Interactions by Assessing Signaling in HEK-293T Cells
Given that CB1R is a Gi -coupled GPCR and its activation leads to inhibit adenylyl cyclase activity and reduce intracellular cAMP levels, we first assessed the cAMP signaling pathway. In HEK-293T cells coexpressing CB1R and NMDAR N1 and N2 subunits, stimulation with the CB1R-selective agonist ACEA led to a ∼40% decrease in forskolin-induced cAMP levels (Figure fig2 A), consistent with canonical CB1R signaling. As expected, NMDA alone had no effect on cAMP, since NMDAR does not engage this pathway. Interestingly, costimulation with ACEA and NMDA did not alter the magnitude of the cAMP response compared to ACEA alone. However, the CB1R-mediated effect was blocked not only by the CB1R antagonist SR141617 (rimonabant) but also by MK-801, a noncompetitive NMDAR antagonist. This effect, known as cross-antagonism, suggests functional coupling between the two receptors and is considered a key signature of receptor–receptor interactions. ref36−ref37 ref38 ref39
To further assess functional crosstalk, we focused on the MAPK signaling pathway that may be engaged upon agonist-induced activation of the CB1R and, also, of the NMDAR. Although both CB1R and NMDAR stimulation induced ERK1/2 phosphorylation (Figure fig2 B), coactivation did not lead to an additive response. ERK1/2 phosphorylation induced by NMDA was inhibited not only by MK-801, but also by rimonabant, and vice versa. Therefore, cross-antagonism was observed across both signaling pathways analyzed.
Cytosolic calcium levels signaling induced by NMDA were determined. In HEK-293T cells coexpressing CB1R and NMDAR N1 and N2 subunits, NMDAR reconstitution occurred and its stimulation produced a robust cytosolic calcium increase. ACEA partially attenuated the NMDA-induced calcium response (Figure fig2 C), indicating a negative crosstalk. Cross-antagonism was again observed in this assay, reinforcing the idea of molecular and functional links between both receptors.
To explore whether cross-modulation extends to CB1R receptor desensitization and internalization, we evaluated β-arrestin-2 recruitment upon receptor activation by ACEA. Using a BRET-based β-arrestin-2 recruitment assay, we found that the agonist of the NMDAR significantly enhanced ACEA-induced β-arrestin-2 binding to CB1R (Figure fig2 D), suggesting that NMDAR activity may contribute to shortening the duration of the effect of cannabinoids acting on CB1R. Again, a cross-antagonism was detected.
Finally, dynamic mass redistribution (DMR) assays were performed to monitor global cellular responses such as receptor activation and cytoskeletal reorganization. The DMR data mirrored the results of the other signaling assays studied, displaying clear patterns of negative crosstalk and cross-antagonism between CB1R and NMDAR (Figure fig2 E).
Altogether, these findings provide strong functional evidence of receptor cross-modulation between CB1R and NMDAR, reinforcing the hypothesis that these receptors form functional complexes with the capacity to modulate each other’s signaling.
Evidence of CB1–NMDA Receptor–Receptor Interactions in Microglia
To investigate the expression of CB1R-N1 subunit complexes in primary microglia from mouse brain, a proximity ligation assay (PLA) was performed. Using this technique, interactions of the two proteins were observed as punctate red clusters (Figure fig3 A,B). The staining was observed in a high percentage of naïve/resting microglial cells (>80%), with an average of seven puncta per positive cell. Furthermore, microglia activated using 1 μM LPS and 200 U/mL IFN-γ showed an important increase in the CB1R-N1 complex expression, with >90% of cells showing red clusters and an average of 18 puncta per positive cell. Primary microglia (naïve/resting) obtained from the brain of the APPSw/Ind transgenic mouse model of AD also showed a high percentage of cells (>80%) expressing clusters with an average of 10 puncta per positive cell (Figure fig3 A,B).
The presence of complexes of CB1R and the N1 subunit of the NMDAR in microglia prompted us to investigate the effect of activating the CB1 and/or the NMDA receptors. First, we measured intracellular cAMP levels in both naïve (resting) and activated (under inflammatory stimuli) microglia from nontransgenic animals. Treatment with the CB1R agonist ACEA significantly attenuated forskolin-induced cAMP elevation, consistent with canonical Gi -mediated signaling. However, this effect was counteracted by NMDA, which prevented the CB1R-mediated reduction in cAMP levels (Figure fig3 C,D). Notably, the cross-antagonism observed in HEK-293T cells was neither found in resting nor activated microglia from nontransgenic animals. In contrast, naïve (resting) microglia derived from the APPSw/Ind mouse model of AD exhibited a fairly distinct pattern because NMDA not longer blocked Gi -mediated signaling upon CB1R activation by ACEA (Figure fig3 G). These differences may reflect variations in heteromer composition or the involvement of additional proteins in the receptor complex under overexpression of the mutant form of human amyloid precursor protein.
Next, we examined the mitogen-activated protein kinase (MAPK) signaling pathway by measuring ERK1/2 phosphorylation. In both naïve (resting) and activated microglia from nontransgenic mice, NMDA blocked CB1R-induced ERK1/2 phosphorylation. In addition cross-antagonism was found as CB1R-mediated effects were inhibited not only by the selective CB1R antagonist rimonabant but also by the NMDAR antagonist MK-801 (Figure fig3 E,F). In primary naïve microglia from APPSw/Ind mice, both ACEA and NMDA led to a significant increase in ERK1/2 phosphorylation. However, cotreatment did not significantly increase ERK1/2 phosphorylation over basal levels. This result and the reciprocal cross-antagonism (Figure fig3 H) are evidence of cross-modulation likely derived from a direct interaction between the two receptors.
Functional CB1-NMDA Receptor–Receptor Crosstalk in Primary Cortical and Hippocampal Neurons from Nontransgenic Mice
To investigate the presence of CB1R-N1 subunit complexes in primary neurons from the cortex and hippocampus, regions particularly vulnerable in AD, PLA was undertaken. Red fluorescent clusters indicative of protein complexes were detected (Figure fig4 A,B). In cortical neurons, more than 20% of cells exhibited red clusters, with an average of ∼2.5 puncta per positive cell. In hippocampal neurons, >30% of cells displayed red puncta, also averaging ∼2.5 puncta per positive cell. These findings align with a higher CB1R expression level in hippocampal compared to cortical neurons. Notably, pretreatment with Aβ1–42 oligomers (500 nM, 48 h) significantly increased both the proportion of neurons displaying CB1R-N1 complexes and the number of puncta per positive cell. Following Aβ1–42 exposure, >40% of cortical neurons and >60% of hippocampal neurons displayed red clusters, with an average of ∼4 puncta per positive cell in both regions. Negative controls lacking the anti-CB1R primary antibody exhibited minimal signal (<10% of cells with red puncta), confirming the specificity of the staining. These results demonstrate the presence of CB1R-N1 complexes in both cortical and hippocampal neurons, with a marked upregulation following Aβ1–42 treatment.
To investigate the functional consequences of CB1R–NMDAR interactions, we first assessed intracellular cAMP levels. In both cortical and hippocampal neurons, stimulation with the CB1R agonist ACEA reduced forskolin-induced cAMP accumulation, consistent with Gi protein coupling. This effect was abolished by coapplication of NMDA, suggesting a negative crosstalk between the receptors (Figure fig4 C,E). Activation of either receptor alone induced ERK1/2 phosphorylation; however, simultaneous stimulation with both agonists failed to produce additive or synergistic effects. In hippocampal neurons, cotreatment with ACEA and NMDA did not enhance ERK1/2 phosphorylation beyond basal levels (Figure fig4 D,F). Cross-antagonism, a hallmark of receptor–receptor interactions, was evident in both signaling pathways in cortical and hippocampal neurons. Antagonism of either receptor disrupted the response mediated by the other, supporting functional interplay between the CB1R and NMDAR systems.
Hippocampal neurons treated with Aβ1‑42 exhibited signaling profiles similar to those of untreated cells, with cross-antagonistic interactions remaining intact (Figure fig4 I,J). Cross-antagonism was consistently observed in all tested neurons, regardless of cortical or hippocampal origin, Aβ1‑42 exposure, or assay type (cAMP or ERK1/2 phosphorylation; Figure fig4 C–J). Furthermore, PLA signals indicative of CB1R–N1 subunit interactions were significantly enhanced following Aβ1‑42 treatment in both cortical and hippocampal neurons, suggesting that the amyloid peptide promotes heteromer formation (Figure fig4 A,B). However, in cortical neurons exposed to Aβ1‑42, NMDA no longer inhibited the CB1R-mediated suppression of forskolin-induced cAMP (Figure fig4 G). This dissociation between receptor complex expression and functional output implies the involvement of additional molecular mechanisms modulating CB1R–NMDAR signaling post-Aβ1‑42 exposure. Notably, in Aβ1‑42-treated cortical neurons, the combined effect of ACEA and NMDA on signaling responses diverged from that observed with ACEA and MK-801, unlike in untreated cells where both treatments yielded similar outcomes (Figure fig4 G,H). Importantly, experiments performed in cortical and hippocampal slices revealed a negative crosstalk, supporting the presence of CB1R–NMDAR heteromers that are functionally interacting in the native brain tissue (Figure fig4 K,L).
ACEA Reduces NMDA-Mediated Neurotoxicity
To assess whether activation of CB1R mitigates neurotoxicity induced by NMDA, neurite formation was evaluated in primary cortical and hippocampal neurons treated with ACEA, NMDA, or a combination of both. Immunocytochemistry was performed using an anti-Nectin antibody to visualize neurite patterning, and an anti-F-actin antibody conjugated to Alexa Fluor 488 to label neurons. The results showed that NMDA treatment significantly reduced neurite formation, while ACEA alone had no detectable effect compared to vehicle-treated controls in both cortical and hippocampal neurons. Interestingly, cotreatment with NMDA and ACEA partially counteracted the neurite loss induced by NMDA, indicating a neuroprotective effect of CB1R activation (Figure fig5 ). To further explore this protective role, the same experimental paradigm was applied to neurons pretreated with Aβ1–42 (500 nM) for 48 h. Aβ1–42 exposure led to a marked reduction in neurite formation, which was further exacerbated by NMDA treatment. Notably, ACEA treatment fully prevented the neurite loss induced by Aβ1–42 and partially reversed the additional effect caused by NMDA (Figure fig5 ). Together, these findings suggest that CB1R activation can protect cortical and hippocampal neurons from neurotoxic insults induced by NMDA and Aβ1–42, likely through mechanisms related to the preservation of neuronal plasticity.
Functional CB1-NMDA Receptor–Receptor Crosstalk in Primary Neurons from an APPSw/Ind Mouse Model
After showing evidence that treatment with Aβ1–42 enhances the expression of CB1R-N1 complexes in primary neurons of nontransgenic mice, PLA allowed to identify a higher amount of CB1R-N1 complexes in neurons from the APPSw/Ind mouse AD model. The percentages of cells expressing these protein complexes were 56 and 73 in cortical and hippocampal neurons, respectively, with a similar number of puncta per positive neuron (ca. 4.5 puncta). Such expression is even higher than that displayed by neurons from nontransgenic mice after the treatment with Aß1–42. cAMP level determination assays were conducted in primary cortical and primary hippocampal neurons of the APPSw/Ind mouse. When analyzing the CB1R–NMDAR functionality in cortical and in hippocampal neurons from APPSw/Ind mice, it was observed that NMDA minimally affected the ACEA-induced decrease of cAMP while cross-antagonism was maintained (Figure fig6 A,B). The cross-antagonism was also detected in the ERK1/2 phosphorylation assays (Figure fig6 C,D). Overall, the results were similar to those obtained in primary neurons of nontransgenic mice treated with Aß1–42.
Discussion
AD is the most common form of dementia, accounting for over 60% of cases worldwide. Despite significant research efforts, its etiology remains unclear, and current treatments are only symptomatic. One such treatment is memantine, an NMDAR antagonist, which underscores the relevance of NMDA receptors. NMDARs exert protective effects when synaptically localized but can be neurotoxic when activated at extrasynaptic sites. ref8,ref40,ref41
Sánchez-Blázquez et al. (2013) proposed that CB1Rs counteract NMDA-mediated excitotoxicity via direct interaction with the N1 subunit of NMDAR, stabilized by histidine triad nucleotide-binding protein 1 (HINT1).ref24 This interaction allows CB1Rs to reduce surface NMDARs through cointernalization, diminishing excitotoxic signaling (e.g., calcium influx, NO production, and zinc mobilization). The physiological function depends on the structural integrity of the complex, which is disrupted by PKA activation or in HINT1-deficient mice. These findings established CB1R as a context-sensitive modulator of glutamatergic toxicity. ref42,ref43 These previous studies focused primarily on neurons and nitric oxide, overlooking microglia, the main producers of NO in the CNS and key players in neuroinflammation and AD. Here, we present the first evidence of CB1R–NMDAR interactions in primary microglia, including in naïve/resting and LPS/IFN-γ-activated states. PLA revealed CB1R-N1 complexes in microglia from nontransgenic mice, with increased expression under inflammatory stimuli. Functional assays showed a bidirectional crosstalk in cAMP and MAPK pathways, highlighting that these interactions are not neuron-specific and are influenced by cellular context.
Previous studies focused primarily on neurons and nitric oxide, overlooking microglia, the main producers of NO in the CNS and key players in neuroinflammation and AD. Here, we present the first evidence of CB1R–NMDAR interactions in primary microglia, including in resting and LPS/IFN-γ-activated states. PLA revealed an increased expression of CB1R–N1 complexes under inflammatory stimuli. Functional assays showed a bidirectional crosstalk in cAMP and MAPK pathways, highlighting that these interactions are not neuron-specific and are influenced by cellular context.
Interestingly, while cross-antagonism between CB1R and NMDAR was consistently observed in HEK-293T cells and primary neurons, it was notably absent in naïve microglia from nontransgenic mice, yet clearly present in microglia derived from APPSw/Ind mice. This finding suggests that CB1R–NMDAR functional interactions are differentially regulated in AD, potentially reflecting disease-driven alterations in microglial activation states. In a previous study, we reported that CB1R expression is upregulated in naïve microglia from APPSw/Ind mice, which exhibited a functional profile resembling that of activated microglia from wild-type animals.ref26 These observations support the idea that, in the AD context, microglia may adopt a chronically activated phenotype, likely aligned with an M2-like, neuroprotective state.
We further confirmed the presence of CB1R–N1 complexes in cortical and hippocampal neurons, with Aβ1–42 treatment enhancing their formation. Despite increased expression, functional crosstalk varied with region, treatment, and genotype, indicating that receptor abundance alone does not determine interaction strength. In cortical neurons, NMDA failed to block ACEA-induced cAMP inhibition after Aβ1–42 exposure, whereas hippocampal neurons retained classical negative crosstalk patterns.
Importantly, the functional outcomes of CB1R–NMDAR interactions are strongly context-dependent, suggesting the involvement of additional proteins within the signalosome that modulate these interactions. This is exemplified by two key observations. First, cross-antagonism, a hallmark of receptor–receptor interaction, was consistently observed across conditions, with one notable exception: naïve microglia from nontransgenic mice, where it was absent. Second, the negative modulation of CB1R G i coupling by NMDA varied by cell type and disease context. This inhibitory effect was detected in hippocampal neurons, regardless of Aβ exposure or transgenic status, but was not observed in cortical neurons under any condition. In microglia, NMDA-mediated disruption of CB1R G i coupling was evident in both resting and activated cells from nontransgenic mice, but this effect was lost in resting microglia from the APPSw/Ind model. These cell-type-specific differences highlight the complexity and plasticity of the CB1R–NMDAR signalosome and point to a dynamic interplay modulated by yet-to-be-identified scaffold or regulatory proteins.
These findings suggest that signaling via the CB1R–NMDAR is shaped by multiple factors: cell type, developmental stage, disease context, and the molecular composition of the signalosome. ref44,ref45 For example, changes in NMDAR subunit composition (e.g., N2 vs N3) or availability of scaffold proteins and kinases (e.g., β-arrestins, CaMKII, and PKA) could modulate receptor conformation and signaling outcomes. ref46−ref47 ref48 Supporting this, binding assays showed reduced ACEA affinity when CB1R was coexpressed with N1 and N2 subunits, suggesting conformational shifts.
Notably, CB1R activation rescued neurite loss induced by NMDA and Aβ1–42, highlighting its protective role. This aligns with the hypothesis that CB1R–NMDAR heteromers act as context-dependent signalosomes capable of integrating extracellular cues with intracellular responses.
Overall, our findings reinforce the therapeutic potential of targeting CB1R–NMDAR interactions in AD and neuroinflammation. ref49−ref50 ref51 Unlike direct NMDAR antagonists, which can disrupt normal synaptic activity, ref52,ref53 CB1R agonists offer a more selective approach by modulating pathological signaling via receptor–receptor interactions. Future therapies might aim to stabilize or bias these complexes using allosteric modulators tailored to preserve physiological glutamatergic function while attenuating excitotoxicity.
Conclusions
This study demonstrates that CB1 and NMDA receptors form functional heteromeric complexes in both neurons and microglia, under physiological conditions and in pathological contexts such as AD models. These interactions are not merely structural but lead to bidirectional signaling crosstalk, notably including cross-antagonism across key intracellular pathways. CB1R–NMDAR complexes do not consistently predict their functional impact, pointing to the critical influence of the surrounding signalosome composition. This suggests that additional, cell-specific molecular components govern the signaling outcomes of these receptor complexes.
Our results show that functional complexes involving CB1 and NMDA receptors are highly plastic and context-dependent, integrating diverse intracellular components that shape their signaling outputs. This complexity explains the cell- and condition-specific effects of cannabinoid signaling and highlights the therapeutic potential of selectively targeting CB1R within such receptor complexes. The ability of CB1R to modulate NMDAR activity indirectly, via receptor–receptor interactions, provides a compelling therapeutic avenue. Targeting CB1R within these defined signalosomes, rather than using global NMDAR antagonists, may allow for selective attenuation of excitotoxicity, preservation of neuronal structure, and modulation of neuroinflammatory processes in AD and related neurodegenerative disorders.
Acknowledgements
I.R.-R. is a Serra Húnter fellow.
Notes
Data can be obtained from the corresponding author, G.N., upon reasonable request.
Notes
Conceptualization: G.N.; data curation: G.N., I.R., J.B.R., J.L., C.P.-O., T.C., and A.C.; formal analysis: A.C., I.R., J.L., I.R.-R.; funding acquisition: G.N. and R.F.; investigation: I.R., J.B.R., J.L., C.P.-O., T.C., E.C., A.C., and G.N.; methodology: A.C., I.R., J.L., I.R.-R., and E.S.; project administration: G.N. and R.F.; resources C.A.S. and E.S.; supervision: G.N., A.C., I.R.-R., and R.F.; validation: G.N., A.C., I.R.-R., and R.F.; visualization: G.N. and J.B.R.; writingoriginal draft: R.F.; and writingreview and editing: C.A.S., A.C., I.R.-R., and R.F. All authors approved the version submitted.
Notes
This work was supported by grants PID2020-113430RB-I00 (G.N.) and PID2022-137668OB-I00 (C.A.S.) funded by Spanish MCIN/AEI/10.13039/501100011033 and, as appropriate, by “ERDF A way of making Europe”, by the “European Union” or by the “European Union Next Generation EU/PRTR”. The research groups of the University of Barcelona and of the Autonomous University of Barcelona are considered of excellence (grup consolidat #2021 SGR00304 and #2021 SGR00142, respectively) by the Regional Catalonian Government, Generalitat de Catalunya. The funding bodies played no role in the study design; the collection, analysis, or interpretation of data; the writing of the manuscript; or the decision to submit the paper for publication.
Notes
Animal handling, sacrifice, and further experiments were conducted according to the guidelines set in Directive 2010/63/EU of the European Parliament and the Council of the European Union that is enforced in Spain by National and Regional organisms; the 3R rule (replace, refine, reduce) for animal experimentation was also considered. By the current legislation, protocol approval is not needed if animals are sacrificed to obtain a specific tissue. All animal experiments have been conducted according to ARRIVE guidelines.
Notes
The authors declare the following competing financial interest(s): Natural compounds derived from Cannabis sativa L. used in the laboratory of Rafael Franco (University of Barcelona) are provided by Phytoplant Research S.L. (Crdoba, Spain). However, the submitted manuscript does not include any data obtained using these compounds. None of the authors receive any salary from Phytoplant Research S.L., nor do they hold shares or have any other conflicting interests with the company.
Glossary
- A2AR
- adenosine A2A receptor
- Aβ
- ß-amyloid
- AD
- Alzheimer’s disease
- APP
- amyloid precursor protein
- AMPA
- α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid
- BRET
- Bioluminescence Energy Transfer
- CB1R
- cannabinoid receptor type 1
- CB2R
- cannabinoid receptor type 2
- D1R
- dopamine D1 receptor
- DMEM
- Dulbecco’s modified Eagle’s medium
- ERK1/2
- extracellular signal-regulated kinase 1/2
- FK
- forskolin
- GPCR
- G protein-coupled receptor
- H3R
- histamine H3 receptor
- HINT1
- histidine triad nucleotide-binding protein 1
- HTRF
- homogeneous time-resolved FRET
- IFN-γ
- interferon-γ
- LPS
- lipopolysaccharide
- MAPK
- mitogen-ativated protein kinase
- NMDA
- N-methyl-d-aspartate
- NMDAR
- NMDA receptor
- Rluc
- Renilla luciferase
- YFP
- yellow fluorescent protein