This study was carried out in accordance with the recommendations of “Directive 2010/63/EU.” The protocol was approved by the “iMM’s institutional Animal Welfare Body – ORBEA-iMM and the National competent authority – DGAV (Direcção Geral de Alimentação e Veterinária).”

SVZ and DG neurospheres were prepared from early postnatal (P1-3) Sprague-Dawley rats. SVZ and DG fragments were dissected out from 450 μm-thick coronal brain slices, digested with 0.05% Trypsin-EDTA (Life Technologies, Carlsbad, CA, United States) in Hank’s balanced saline solution (HBSS, Life Technologies), and mechanically dissociated with a P1000 pipette. The originated cell suspension was then diluted in serum-free medium (SFM), composed of Dulbecco’s modified Eagle’s medium/Ham’s F-12 medium with GlutaMAX (DMEM + GlutaMAX, Life Technologies) supplemented with 100 U/mL penicillin and 100 μg/mL streptomycin (Pen/Strep; Life Technologies), 1% B27 (Life Technologies) and growth factors (for SVZ cells: 20 ng/mL EGF; Life Technologies); for DG cells: 20 ng/mL EGF (Life Technologies) and 10 ng/mL FGF-2 (Life Technologies) (proliferative conditions). Single cells were then plated on uncoated Petri dishes at a density of 5,000 cells per cm². SVZ cells were allowed to develop for six days, whereas DG cells were allowed to develop for twelve days, both in a 95% air-5% CO₂ humified atmosphere at 37°C. Rat SVZ and DG neurospheres require different culture conditions, time length and growth factors, because DG-derived neurospheres need more time to reach optimal dimensions when comparing with SVZ (Guo et al., 2012) and SVZ cells do not require the growth factor FGF-2 to expand into neurospheres (Palmer et al., 1995; Kuhn et al., 1997; Woodbury and Ikezu, 2014). Six-day-old SVZ neurospheres and 12-day-old DG neurospheres were adhered for 24 h onto glass coverslips (at a density of approximately 60 neurospheres per well) coated with 0.1 mg/mL poly-D-lysine (PDL, Sigma–Aldrich, St. Louis, MO, United States) in SFM devoid of growth factors (differentiative conditions). One day after plating, the medium was renewed with or without (control) a range of pharmacological treatments for CB₁R and CB₂R ligands, chosen based on their affinity for CB₁R and CB₂R represented by K_i value (Table 1).

To study cell proliferation, plated neurospheres were allowed to develop for 24 h (DIV 1) in the absence (control) or presence of CB₁R and/or CB₂R ligands (Table 1). Neuronal differentiation was assessed by allowing neurospheres to develop for 7 days (DIV 7) in the absence (control) or presence of the aforementioned ligands and an immunocytochemistry for different neuronal markers was performed. To study neuritogenesis, SVZ and DG neurospheres were dissociated using phosphate-buffered saline (PBS) without Mg²⁺/Ca²⁺ and with EDTA (2.5 mM KCl, 1.5 mM KH₂PO₄, 135 mM NaCl, 8 mM Na₂HPO₄, 0.5 mM EDTA.4Na) and plated in differentiative conditions at a density of 10000 cell/cm². DG and SVZ cells were allowed to grow for 2 (DIV 2) and 3 days (DIV 3), respectively, in the absence (control) or presence of the aforementioned ligands and were then fixed and stained for βIII tubulin (neuron-specific class III beta-tubulin).

Whenever cultures needed to be co-treated with both agonists, SVZ or DG cells were primarily treated with CB₁R (ACEA) selective agonist for 30 min prior to CB₂R selective agonist (HU-308) treatment and then grown for 24 h or 7 days in the presence of the ligands (Supplementary Figure S1A). Similarly, treatment with the selective antagonist for CB₁R (AM251) or CB₂R (AM630) was performed for 30 min prior to the treatment with the non-selective cannabinoid receptor agonist WIN55,212-2 or CB₁R (ACEA) or CB₂R (HU-308) selective agonists and then co-incubated for further 24 h or 7 days (Supplementary Figure S1B). Additionally, whenever cultures needed to be treated with a combination of both antagonists and both agonists, SVZ and DG cells were firstly treated with selective antagonists for CB₁R (AM251) and CB₂R (AM630) for 30 min prior to CB₁R selective agonist (ACEA), incubated for further 30 min before CB₂R selective agonist (HU-308) treatment. Thereafter cells were allowed to grow for 24 h in the case of cell proliferation studies or 7 days in the case of neuronal differentiation studies (Supplementary Figure S1C).

To investigate the effect of the different pharmacological treatments on cell proliferation, SVZ and DG cells were exposed to 10 μM 5-bromo-2′-deoxyuridine (BrdU) (Sigma–Aldrich) for the last 4 h of each specific pharmacological treatment (24 h, DIV1). BrdU is a synthetic thymidine analog able to substitute thymidine in the DNA double chain synthesis occurring in dividing cells. Furthermore, neuronal differentiation of proliferating progenitors was assessed in SVZ and DG cells that were incubated in the absence (control) or in the presence of CB₁R/CB₂R ligands together with 10 μM BrdU for 24 h. Then the cells were washed and incubated without BrdU for more 6 days, fixed and co-stained for BrdU and a marker of mature neurons (neuronal nuclear protein, NeuN). For that, SVZ and DG cells were fixed in 4% paraformaldehyde (PFA), BrdU was unmasked by permeabilizing cells in PBS 1% Triton X-100 (Sigma–Aldrich) and DNA was denaturated in 1M HCl. Following blocking in PBS with 0.5% Triton X-100 and 3% bovine serum albumin (BSA), cells were incubated overnight with the anti-BrdU antibody and anti-NeuN antibodies (Table 2). Cells were incubated with the secondary anti-rat Alexa Fluor 488 and anti-rabbit Alexa Fluor 568 (Table 2). Nuclei were stained with Hoechst 33342 (6 μg/mL in PBS, Life Technologies). The final preparations were mounted using Mowiol fluorescent medium.

SVZ and DG cells treated for 7 days (DIV 7) were fixed in 4% PFA, permeabilized and blocked with 0.5% Triton X-100 and 6% BSA in PBS. Cells were then incubated overnight at 4°C with the primary antibodies against different neuronal markers in 0.1% Triton X-100 and 0.3% BSA (w/v) in PBS, and then with the appropriate secondary antibodies in PBS (Table 2). Nuclei counterstaining and mounting were performed as described previously.

Western blotting analysis of CB₁R and CB₂R was performed using SVZ and DG neurospheres that were plated and allowed to develop for 24 h (representing DIV 1) or 7 days (representing DIV 7) in SFM devoid of growth factors. The cells were harvested in RIPA lysis buffer [50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 5 mM ethylenediamine, tetra-acetic acid (EDTA), 0.1%SDS and 1%Triton X-100, containing a protease inhibitor cocktail (pH 7.4, 4°C, Roche, Penzberg, Germany)]. Protein concentration was measured by the Lowry method and samples were treated with SDS-PAGE sample buffer [5x concentrated: 350 mM Tris, 10% (w/v) SDS, 30% (v/v) glycerol, 0.6M DTT, 0.012% (w/v) bromophenol blue], boiled 5 min at 95°C and processed for Western blotting analysis. All samples were applied with same amount of total protein and protein running was performed in SDS-PAGE gels (10% acrylamide/bisacrylamide for resolving and 5% for stacking gels), transferred to polyvinylidene difluoride (PVDF) membranes (GE Healthcare, Buckinghamshire, United Kingdom). Incubations with the primary antibodies against CB₁R (1:500, Frontier Institute Co., Ltd, Japan) and CB₂R (1:1000, Santa Cruz Biotechnology) were performed overnight at 4°C. Membranes were incubated with the respective secondary antibodies: goat IgG anti-guinea pig and goat IgG anti-mouse conjugated with horseradish peroxidase (HRP) (1:10000; Santa Cruz Biotechnology). For endogenous control of immunolabeling, PVDF membranes were reprobed with the antibody against actin (1:1000; #A2006, Sigma–Aldrich). Finally, immunoreactivity was visualized using ECL chemiluminescence detection system (Amersham-ECL Western Blotting Detection Reagents from GE Healthcare, Buckinghamshire, United Kingdom).

CB₂Rs were immunoprecipitated from culture lysates using an antibody-bead mixture previously prepared. Rec-Protein G-Sepharose 4B Conjugate beads (Thermo Fisher Scientific) were first incubated with PBS 0.1% BSA for 1h at 4°C. After washing with PBS and centrifugation at 5000 rpm for 3 min, the pellet composed of beads was incubated for 4 h, at 4°C, with 1 μg of CB₂R antibody. Following several washes, 35 μL of antibody-bead mixture was incubated with 250 μg of total protein (approximately 400 μL of culture lysate) overnight at 4°C. The mixture was centrifuged and the supernatant was collected. The remaining pellet of beads was washed three times with PBS 0.01% Tween 20 and re-suspended and denatured in 35 μL of SDS-PAGE sample buffer. Samples were boiled for 5 min at 95°C. Detection of CB₂R and CB₁R were performed through western blotting, as described above.

To determine the functional neuronal differentiation of SVZ cells, the variations of intracellular calcium concentration ([Ca²⁺]_i) in single cells following stimulation with 50 mM KCl and 100 mM histamine (Sigma–Aldrich) were analyzed as previously described for SVZ cells (Agasse et al., 2008). KCl depolarization causes an increase in [Ca²⁺]_i in neurons, whereas stimulation with histamine leads to an increase in [Ca²⁺]_i in stem/progenitor cells (Agasse et al., 2008; Xapelli et al., 2013). For SCCI analysis SVZ cultures plated in PDL-coated glass bottom cell culture dishes (Nest, NJ, United States) and grown for 7 days were loaded for 45 min with 5 μM Fura-2/AM (Invitrogen) and 0.02% pluronic acid F-127 (Invitrogen) in Krebs solution (132 mM NaCl, 4 mM KCl, 1.4 mM MgCl₂, 1 mM CaCl₂, 6 mM glucose, 10 mM HEPES, pH 7.4), in an incubator with 5% CO₂ and 95% atmospheric air at 37°C. After a 5 min postloading period at RT in Krebs solution without Fura-2/AM and pluronic acid, to obtain a complete hydrolysis of the probe, the dish was mounted on an inverted microscope with epifluorescent optics (Axiovert 135TV, Zeiss) equipped with a xenon lamp and band-pass filters of 340 and 380 nm wavelengths. Cells were continuously perfused with Krebs solution and stimulated at defined periods of time by applying high-potassium Krebs solution (containing 50 mM KCl, isosmotic substitution with NaCl) and 100 μM histamine (Supplementary Figure S2). Throughout the experiments, the cells were continuously superfused at 1.5 mL/min with physiological solution. KCl and histamine were applied focally through a drug filled micropipette placed under visual guidance. Image pairs obtained every 200 ms by exciting the preparations at 340 and 380 nm were taken to obtain ratio images. Excitation wavelengths were changed through a high-speed wavelength switcher, Lambda DG-4 (Sutter Instrument, Novato, CA, United States), and the emission wavelength was set to 510 nm. Image data was recorded with a cooled CCD camera (Photometrics CoolSNAP fx) and processed and analyzed using the software MetaFluor (Universal Imaging, West Chester, PA, United States). Regions of interest were defined manually over the cell profile. KCl and histamine peaks given by the normalized ratios of fluorescence at 340/380 nm, at the proper time periods, were used to calculate the ratios of the responses.

Fluorescence images were recorded using an LSM 880 confocal microscope or an Axiovert 200 inverted widefield fluorescence microscope (both from Carl Zeiss Inc., Göttingen, Germany), with a 40x objective. Images were recorded using the softwares ZEN 2.1 (black edition) or AxioVision 4 (both from Carl Zeiss Inc.).

In all experiments, measurements were performed at the border of SVZ and DG neurospheres, where migrating cells form a pseudo-monolayer of cells. In every independent experiment, which corresponds to one independent neurosphere culture from one litter, each condition was measured in three different coverslips. Percentages of BrdU, BrdU/NeuN, NeuN, DCX and DCX/NeuN immunoreactive cells were calculated from cell counts of five independent microscopic fields in each triplicated coverslip (representing n = 1) with a 40x objective (∼200–400 cells per field). For quantification of neuritogenesis neurites were manually traced using NeuronJ v1.4.3, a plugin from ImageJ v1.46r (10 random images were captured per glass coverslip, in triplicates), measuring the total neurite length, the maximal neurite length, as well as the number of primary dendrites and branch points. An average of at least 10 neurons were acquired per glass coverslip for each experiment. All experiments were analyzed in a double-blind fashion and part of data was normalized to each corresponding control.

For SCCI experiments, the percentages of neuronal-like responding cells (with a Hist/KCl ratio below 0.8) and of immature-like responding cells (with a Hist/KCl ratio between 1 and 1.3) were calculated on the basis of one microscopic field per glass coverslip, containing approximately 100 cells, in a total of 3–4 independent cultures where each condition are duplicate.

Data are expressed as mean ± standard error of the mean (SEM). Statistical significance was determined using one-way analysis of variance followed by Sidak’s or Dunnett’s-multiple comparison test, with p < 0.05 considered to represent statistical significance.

Using neurospheres as a method to study postnatal neurogenesis it was possible to observe that SVZ neurospheres are composed by stem/progenitor cells since they are formed by undifferentiated cells that express Sox2 (transcription factor important to maintain self-renewal) and Nestin (intermediate filament protein expressed by neuronal precursor cells) and, incorporate BrdU (Figure 1A). Importantly plating the neurospheres in differentiative conditions for 7 days (DIV 7) is sufficient to induce neuronal differentiation since a number of SVZ-derived cells express neuronal immature markers, such as doublecortin (DCX) and βIII tubulin and, also a marker for mature neurons, such as NeuN (Figure 1C). Furthermore, immunolabeling for the vesicular GABA transporter (VGAT, marker for GABAergic neurons) and the tyrosine hydroxylase (TH, marker for dopaminergic neurons) showed that SVZ-derived neurons express these proteins at DIV 7 (Figures 1E,F), which is in accordance with the fact that the two major phenotypes of SVZ-derived neurons are GABAergic and dopaminergic neurons. Similarly to SVZ cultures, DG neurospheres are also composed by undifferentiated cells expressing the stem/progenitor cell markers Sox2 and Nestin and with the ability to incorporate BrdU (Figure 1B). We have also observed that some DG-derived cells after 7 days in differentiative conditions (DIV 7) also express the immature markers DCX and βIII tubulin and the mature marker NeuN (Figure 1D). Additionally, DG-derived neurons at DIV 7 were found to express VGAT and vesicular glutamate transporter (VGlut1, marker for glutamatergic neurons), which reflects the two major phenotypes of DG-derived neurons, i.e., GABAergic and glutamatergic neurons (Figures 1G,H). Importantly, the expression of CB₁R and CB₂R in SVZ- and DG-derived cells was observed by immunocytochemistry and by western blotting (Figures 1I–K). In fact, CB₁R and CB₂R expression was found in both DIV 1 and DIV 7 SVZ and DG cells as well as in adult tissue (Figure 1K). In particular, a band for CB₁R was found in the predicted molecular weight (52 kDa) and other bands at higher molecular weights (between 52 and ∼70 kDa) were also found, which probably correspond to glycosylated forms of the receptor (Song and Howlett, 1995; Bermúdez-Silva et al., 2008). Similarly, a band for CB₂R was found at the predicted molecular weight (∼35 kDa) and other at ∼60 kDa which may also represent a glycosylated form (Gong et al., 2006; Bermúdez-Silva et al., 2008).

To determine if cannabinoid receptor activation modulates SVZ cell proliferation, increasing concentrations (100 nM, 300 nM, 1 μM) of selective CB₁R (ACEA) and CB₂R (HU-308) agonists and, non-selective CB₁R and CB₂R receptor agonist (WIN55,212-2) were applied on SVZ cells in SFM devoid of growth factors for 24 h. BrdU, a thymidine nucleotide analog, was added during the last 4 h of culture to label cells that went through S-phase. After fixation, incorporated BrdU was immunolabeled and positive nuclei were counted (Figure 2A). It was observed a significant increase in the number of BrdU-positive cells when cultures were incubated with 1 μM of the CB₁R selective agonist ACEA, while 100 or 300 nM did not promote cell proliferation (control: 4.04 ± 0.18%; ACEA 1 μM: 5.67 ± 0.56%; n = 3, p < 0.05) (Figures 2B,C). No significant changes were observed in the number of BrdU-immunopositive nuclei obtained in cultures incubated with all the aforementioned concentrations of either the CB₂R selective agonist HU-308 (Figures 2D,E) or the non-selective CB₁R and CB₂R agonist WIN55,212-2 compared with control cultures (Figures 2F,G). Therefore, concentrations of 1 μM were chosen based on the concentration-response curve.

We then investigated the interaction between CB₁R and CB₂R by combining selective activation of both CB₁R and CB₂R agonists. As previously described, it was detected a significant increase in the number of BrdU-positive cells when cultures were incubated with the CB₁R selective agonist ACEA (control: 100.01 ± 0.15%; ACEA 1 μM: 136.50 ± 5.15%; n = 11, p < 0.001) compared with control cultures (Figures 3A,B). Concomitantly, no significant changes were obtained in cultures incubated with the CB₂R selective agonist HU-308. Surprisingly when both CB₁R and CB₂R selective agonists were co-incubated the increase in cell proliferation mediated by CB₁R activation was inhibited and therefore, had an effect similar to the observed by incubating with the non-selective CB₁R and CB₂R agonist WIN55,212 (Figures 3A,B). This suggests a role for CB₂R in modulating the stimulatory effect promoted by CB₁R on SVZ cell proliferation. To further elucidate the role of this CB₁R-CB₂R crosstalk on SVZ cell proliferation, cells were co-incubated with selective CB₁R and CB₂R selective antagonists (AM251 and AM630, respectively). The effect promoted by CB₁R selective activation was blocked either in the presence of the CB₁R selective antagonist AM251 (ACEA 1 μM + AM251 1 μM: 71.91 ± 12.49%; n = 3, p < 0.001) or in the presence of the CB₂R selective antagonist AM630 (ACEA 1 μM + AM630 1 μM: 99.36 ± 7.76%; n = 5, p < 0.05) (Figure 3C), suggesting a putative interaction between CB₁R and CB₂R. Moreover, cells were further co-incubated with both CB₁R and CB₂R selective agonists (ACEA + HU-308) while being also incubated with either a CB₁R or a CB₂R antagonist, the data obtained being summarized in Figure 3D. When both selective agonists were present (ACEA + HU-308) and also in the presence of the CB₂R selective antagonist, AM630, the effect promoted by ACEA alone on SVZ cell proliferation was rescued (ACEA 1 μM: 136.50 ± 5.15%; ACEA 1 μM+HU-308 1 μM + AM630 1 μM: 138.10 ± 4.77%; n = 5, p < 0.05). Incubation with the CB₁R selective antagonist, AM251, prior to co-exposure with both selective agonists (ACEA + HU-308) had no effect. Moreover, when cells were co-incubated with both CB₁R and CB₂R selective agonists and antagonists no significant changes were observed in the number of BrdU-positive cells (control: 100.00 ± 0.05%; ACEA 1 μM + HU-308 1 μM + AM251 1 μM + AM630 1 μM: 92.40 ± 18.09%; n = 3) (Figure 3E). We also did not observe any significant alterations when incubating cultures with both selective antagonists alone (Figures 3C–E).

Taken together, the above results indicate that the effect observed on SVZ cell proliferation is via CB₁R and that it is markedly affected while perturbing the degree of activation of CB₂R. Overactivation of CB₂R by exogenous agonists hampers the action of CB₁R, but full blockade of CB₂R in the absence of exogenous CB₂R agonist also prevents the influence of CB₁R upon SVZ cell proliferation.

To disclose whether cannabinoid receptor activation affects SVZ neuronal differentiation, SVZ cells were treated with increasing concentrations (100 nM, 300 nM, 1 μM) of selective CB₁R (ACEA) and CB₂R agonists (HU-308) and non-selective CB₁R and CB₂R agonist (WIN55,212-2) for 7 days in SFM without growth factor (Figure 4A). A significant increase in the number of NeuN-positive cells was observed when SVZ cultures were incubated with ACEA 1 μM but not with 100 or 300 nM, (control: 1.96 ± 0.14%; ACEA 1 μM: 3.72 ± 0.40%; n = 3–4, p < 0.05) (Figures 4B,C). When incubating SVZ cultures with HU-308, the CB₂R selective agonist, we observed a significant increase in the number of NeuN-positive cells in all tested concentrations [control: 1.96 ± 0.14%; HU-308: 3.52 ± 0.27% (100 nM), 3.28 ± 0.38% (300 nM), 3.75 ± 0.32% (1 μM); n = 3–4, p < 0.05 and p < 0.01] (Figures 4D,E). Moreover, it was observed a marked increase in the number of NeuN-immunopositive nuclei when cells were incubated with increasing concentrations of WIN55,212-2, which reached statistical significance for concentrations ≥300 nM [control: 2.02 ± 0.28%; WIN55,212-2: 4.17 ± 1.13% (100 nM), 4.37 ± 1.23% (300 nM), 3.83 ± 0.47% (1 μM); n = 5–12, p < 0.05) (Figures 4F,G]. As with cell proliferation, 1 μM was the concentration selected for the following experiments. Interestingly, when we assessed whether selective or non-selective cannabinoid receptor activation would have an impact in the number of immature neurons, we observed a significant and robust increase in the number of DCX-positive cells with all receptor agonists used (control: 26.55 ± 2.12%; ACEA 1 μM: 51.43 ± 1.06%; HU-308 1 μM: 50.57 ± 2.75%; WIN55,212-2 1 μM: 49.30 ± 5.56%; n = 3, p < 0.01 and p < 0.001) (Figures 4H,I). To further support the role of cannabinoids in SVZ neuronal differentiation, we used a BrdU pulse for the first 24 h of treatments (CB₁R and CB₂R selective and non-selective activation) followed by a chase of 6 days (without BrdU) in the absence (control) or in the presence of the drug treatments (Figure 4A). We observed a 2.5-fold increase in the number of BrdU/NeuN-positive cells in all treated conditions (control: 5.97 ± 0.95%; ACEA 1 μM: 15.70 ± 1.38%; HU-308 1 μM: 15.40 ± 0.87%; WIN55,212-2 1 μM: 16.10 ± 0.49%; n = 4; p < 0.001), indicating that cannabinoid treatment is interfering in the early stages of SVZ neuronal differentiation, committing progenitors toward a neuronal fate (Figures 4J,K).

To discriminate between the relative role of CB₁R and CB₂R upon neuronal differentiation, selective agonists for CB₁R (ACEA, 1 μM) and for CB₂R (HU-308, 1 μM) were used. Treatment of SVZ cells with either ACEA 1 μM or with HU-308 1 μM induced a significant (p < 0.001, n = 11–14) increase in the number of NeuN-positive cells (control: 99.99 ± 0.03%; ACEA 1 μM: 157.10 ± 8.50%; HU-308 1 μM: 143.00 ± 7.62%) (Figures 5A,B). The facilitatory effect upon neuronal differentiation was still evident when both agonists were added together (ACEA 1 μM + HU-308 1 μM: 135.20 ± 14.23%), but remarkably, the effect of both agonists was not additive, with a similar effect comparing with the non-selective CB₁R and CB₂R WIN55,212-2 (Figures 5A,B). The effect caused by CB₁R selective activation was blocked either in the presence of the CB₁R selective antagonist AM251 (ACEA 1 μM + AM251 1 μM: 94.18 ± 6.17%; n = 5–14, p < 0.001) or in the presence of the CB₂R selective antagonist AM630 (ACEA 1 μM + AM630 1 μM: 91.74 ± 11.10%; n = 5, p < 0.001) (Figure 5C). Similarly, the effect caused by CB₂R activation was blocked either in the presence of the CB₁R selective antagonist AM251 (HU-308 1μM + AM251 1 μM: 105.20 ± 9.02%; n = 5–15, p < 0.01) or in the presence of the CB₂R selective antagonist AM630 (HU-308 1 μM + AM630 1 μM: 105.20 ± 12.48%; n = 5, p < 0.01) (Figure 5D). This crossed-antagonism strongly suggests the existence of a CB₁R-CB₂R interaction to regulate SVZ neuronal differentiation. To further confirm the need of an interaction between CB₁R and CB₂R to modulate SVZ neuronal differentiation, co-incubation of both selective agonists and antagonists was performed, the results obtained being depicted in Figures 5E,F. Incubation of the CB₁R selective antagonist, AM251, together with both selective receptor agonists (ACEA + HU-308) induced no significant changes in the number of NeuN-positive cells when compared to control conditions (control: 100.02 ± 0.67%; ACEA 1 μM + HU-308 1 μM + AM251 1 μM: 117.40 ± 8.13%; n = 3–12) (Figure 5E). Interestingly, incubation of the CB₂R selective antagonist, AM630, with both selective agonists allowed an increase in the number of NeuN-positive cells (ACEA 1 μM + HU-308 1 μM + AM630 1 μM: 151.10 ± 10.47%; n = 6, p < 0.05) (Figure 5E), indicating that the influence of CB₁R is preponderant over CB₂R with respect to the modulation of SVZ neuronal differentiation. As expected, the effect promoted by both selective agonists, when added together, was abolished in the presence of both receptor antagonists (Figure 5F) (ACEA 1 μM + HU-308 1 μM + AM251 1 μM + AM630 1 μM: 103.50 ± 7.91%; n = 3–12). Additionally, no significant alterations were found when incubating cultures with both selective antagonists alone (Figures 5C–F). Interestingly, the effect promoted by non-selective activation of CB₁R and CB₂R with WIN55,212-2 on SVZ neuronal differentiation was blocked when cells were co-incubated with the CB₁R selective antagonist AM251 (WIN55,212-2 1 μM + AM251 1 μM: 89.23 ± 9.60%) (Figure 5G) or with the CB₂R selective antagonist AM630 (WIN55,212-2 1 μM + AM630 1 μM: 91.43 ± 16.07%) (Figure 5H), showing that the effect of WIN 55,212-2 on SVZ cell proliferation is dependent on both CB₁R and CB₂R.

In order to functionally evaluate neuronal differentiation of SVZ cells we used SCCI, as previously described (Xapelli et al., 2013). Briefly, variations of [Ca²⁺]_i at single cell level upon KCl and histamine (Hist) stimulations were measured and Hist/KCl ratios were calculated, therefore reflecting the phenotype of the analyzed cells. Ratios below 0.8 are characteristic of SVZ-derived neuronal-like cells whereas ratios between 1 and 1.3 are characteristic of SVZ-derived immature cells (Agasse et al., 2008). Quantification of the percentage of cells displaying a Hist/KCl ratio below 0.8 showed a concentration-dependent increase in the % of neuronal-like cells with the CB₁R selective agonist ACEA [control: 10.21 ± 1.38%; ACEA: 15.83 ± 7.33% (100 nM), 22.67 ± 2.83% (300 nM), 29.75 ± 5.26% (1 μM); n = 3–4, p < 0.05], the CB₂R selective agonist HU-308 [control: 10.21 ± 1.38%; HU-308: 34.50 ± 3.75% (100 nM), 23.50 ± 6.53% (300 nM), 33.50 ± 5.86 (1 μM); n = 3–4, p < 0.05] and with the CB₁R and CB₂R non-selective agonist WIN55,212-2 [control: 10.21 ± 1.38%; WIN55,212-2: 20.17 ± 5.05% (100 nM), 23.67 ± 4.88% (300 nM), 32.05 ± 4.93% (1 μM); n = 3–4, p < 0.05] (Figures 6A,B). Furthermore, selective activation of both receptors (ACEA + HU-308) also promoted a significant increase in the % of neuronal-like cells (ACEA 1 μM + HU-308 1 μM: 36.67 ± 5.40%; n = 3, p < 0.001) (Figures 6A,B). In fact, a three-fold increase in the number of neuronal-like cells resulting from incubation of these cannabinoids receptor agonists at concentrations of 1 μM was observed, indicating a robust cannabinoid-mediated augmentation of neuronal differentiation. Moreover, CB₁R and/or CB₂R selective and non-selective activation were also found to promote a significant decrease in the % of immature-like cells (ratio Hist/KCl 1–1.3) at 1 μM concentration (control: 33.80 ± 5.06%; ACEA 1 μM: 15.25 ± 3.19%; HU-308 1 μM: 8.87 ± 3.16%; WIN55,212-2 1 μM: 14.50 ± 5.65%; ACEA 1 μM+HU-308 1 μM: 10.50 ± 3.68%; n = 3–4, p < 0.05 and p < 0.01) (Figures 6A,C). While control cultures displayed a predominant immature-like profile, characterized by an increase in [Ca²⁺]_i in response to histamine but a small response or no response to KCl stimulation, the majority of ACEA-, HU-308-, WIN55,212-2-treated SVZ cultures displayed an increase in [Ca²⁺]_i in response to KCl but not to histamine stimulation which is consistent with a neuronal-like profile (Figure 6D and Supplementary Figure S2).

Finally, in agreement with the observed proneurogenic action of cannabinoids, we investigated whether our treatments triggered neuronal morphological changes, consistent with neuronal maturation, by looking at neurite outgrowth and branching processes of neurons stained for βIII tubulin (Figures 7A,B). When comparing to control neurons (Figure 7C), incubation of SVZ cultures with ACEA, HU-308 and WIN55,212-2 induced a significant increase in the total length of neurites per neuronal cell (control: 100.0 ± 0.0%; ACEA 1 μM: 124.2 ± 7.60%; HU-308 1 μM: 136.7 ± 6.94%; WIN55,212-2 1 μM: 142.4 ± 11.13%; n = 4–9, p < 0.05, p < 0.01 and p < 0.001) (Figure 7D) and in the length of the maximal neurite (control: 100.0 ± 0.0%; ACEA 1 μM: 113.5 ± 8.66%; HU-308 1 μM: 124.0 ± 5.48%; WIN55,212-2 1 μM: 137.1 ± 5.47%; n = 4–9, p < 0.01 and p < 0.001) (Figure 7F). Furthermore, cannabinoid receptor agonists treatments promoted a marked increase in the number of branch points per cell when compared to control cultures (control: 100.0 ± 0.0%; ACEA 1 μM: 145.3 ± 6.32%; HU-308 1 μM: 168.1 ± 21.26%; WIN55,212-2 1 μM: 161.2 ± 7.22%; n = 4–9, p < 0.01 and p < 0.001) (Figure 7G) while not perturbing the number of primary neurites (Figure 7E).

It was next investigated the effect of cannabinoid receptor activation on DG cell proliferation. For that purpose, DG cells were treated with increasing concentrations (100 nM, 300 nM, 1 μM) of CB₁R and CB₂R selective agonists and a non-selective CB₁R and CB₂R agonist for 24 h in SFM devoid of growth factors (Figure 8A). As shown in Figures 8B–E, CB₁R selective activation with ACEA or CB₂R selective activation with HU-308 did not promote any changes in the number of BrdU-positive cells in any tested concentration when compared with control conditions. Non-selective receptor activation with WIN55,212-2 caused a concentration-dependent increase in BrdU-positive cells, though the effect only reached statistical significance at 1 μM (control: 8.72 ± 3.16%; WIN55,212-2 1 μM: 18.55 ± 2.50%; n = 3–5, p < 0.05) (Figures 8F,G).

To assess the relative role of each cannabinoid receptor, DG cultures were co-incubated with the selective receptor antagonists and agonists. There were no significant changes when incubating cultures with only ACEA or HU-308 (control: 100.00 ± 0.03%; ACEA 1 μM: 109.00 ± 9.65%; HU-308 1 μM: 98.68 ± 4.34%; n = 15, p > 0.05) (Figures 9A,B), when compared to control cultures, concomitant with the abovementioned results. However, when co-incubating cultures with both selective agonists for CB₁R and CB₂R (ACEA + HU-308) a surprisingly significant increase was observed in the number of BrdU-positive cells (ACEA 1 μM + HU-308 1 μM: 162.50 ± 18.83%; n = 10, p < 0.001), similar to the effect observed when cells were incubated with the non-selective receptor agonist WIN55,212-2 (Figures 9A,B). These results suggest that both receptors have to be active to induce DG proliferation. Furthermore, it was observed that in the presence of the CB₁R selective antagonist, AM251, the effect promoted by incubation with both selective receptor agonists was lost (ACEA 1 μM + HU-308 1 μM + AM251 1 μM: 77.95 ± 4.12%; n = 5, p < 0.01) (Figure 9C), whereas co-incubation with the CB₂R selective antagonist AM630 did not block the effect mediated by CB₁R and CB₂R co-activation (ACEA 1 μM + HU-308 1 μM + AM630 1 μM: 162.60 ± 18.44%; n = 5, p < 0.05) (Figure 9C). These data suggest that CB₁R has a leading role in modulating DG proliferation. As could be expected, the effect promoted by co-incubation with both CB₁R and CB₂R selective agonists was completely lost when both receptor antagonists were present together (ACEA 1 μM + HU-308 1 μM + AM251 1 μM + AM630 1 μM: 68.76 ± 10.05%; n = 5, p < 0.01) (Figure 9D). Interestingly, when cultures were incubated with the CB₁R and CB₂R selective antagonists in the absence of agonists, a significant decrease on DG cell proliferation was observed (AM251 1 μM + AM630 1 μM: 60.15 ± 12.00%; n = 5, p < 0.05) (Figure 9D), suggesting a pivotal influence of endocannabinoids upon DG cell proliferation; single incubation with each of the receptor antagonist had no substantial effect (Figures 9C,D). Moreover, the increase on DG cell proliferation promoted by WIN55,212-2 1 μM was blocked either when cells were co-incubated with the CB₁R selective antagonist AM251 (WIN55,212-2 1 μM+AM251 1 μM: 90.93 ± 8.10%; n = 5–23, p < 0.05) (Figure 9E) or with the CB₂R selective antagonist AM630 (WIN55,212-2 1 μM + AM630 1 μM: 96.85 ± 10.86%; n = 5–23, p < 0.05) (Figure 9F), suggesting a CB₁R-CB₂R crosstalk mechanism mediating DG cell proliferation.

To determine the effect of cannabinoid receptor activation on DG neuronal differentiation, DG cells were treated with increasing concentrations (100 nM, 300 nM, 1 μM) of CB₁R and CB₂R selective agonists and a non-selective CB₁R and CB₂R agonist in SFM devoid of growth factors for 7 days (Figure 10A). CB₁R selective activation with ACEA promoted a concentration-dependent increase in the number of NeuN-positive cells, reaching statistical significance at the concentration of 1 μM (control: 2.18 ± 0.33%; ACEA 1 μM: 4.51 ± 0.47%; n = 3–6, p < 0.05) (Figures 10B,C). Similarly, when cultures were incubated with CB₂R selective agonist HU-308 the same concentration-dependent profile was observed (control: 2.43 ± 0.44%; HU-308 1 μM: 4.57 ± 0.46%; n = 3–6, p < 0.05) (Figures 10D,E). An increase in the number of NeuN-positive cells was observed with all concentrations used for the non-selective receptor agonist WIN55,212-2, only reaching statistical significance at the concentration of 1 μM (control: 3.61 ± 0.58%; WIN55,212-2 1 μM: 7.03 ± 1.06%; n = 3–10, p < 0.05) (Figures 10F,G). When assessing the overall influence of cannabinoid receptors on early stages of DG neuronal differentiation by selectively activating CB₁R (ACEA 1 μM) or CB₂R (HU-308 1 μM) or, by non-selective receptor activation (WIN55,212-2 1 μM) we observed an increase in the number of immature neurons characterized by a significant increase in the number of DCX-positive cells (control: 10.73 ± 0.19%; ACEA 1 μM: 17.34 ± 1.13%; HU-308 1 μM: 17.00 ± 0.84%; WIN55,212-2 1 μM: 16.94 ± 1.18%; n = 3, p < 0.01) (Figures 10H,I), showing that activation of cannabinoid receptors functions as an early neurogenic propeller. Similarly to SVZ, we next investigated whether CB₁R and CB₂R activation could be committing DG early progenitors toward a neuronal fate. For that a BrdU pulse was done in the first 24 h of a 7-day treatment with ACEA, HU-308 and WIN55,212-2 (Figure 10A). We observed a substantial increase in the number of BrdU/NeuN-positive cells in all treated conditions (control: 11.55 ± 0.97%; ACEA 1 μM: 27.80 ± 1.69%; HU-308 1 μM: 23.99 ± 2.13%; WIN55,212-2 1 μM: 25.88 ± 4.76%; n = 3, p < 0.05 and p < 0.01) (Figures 10J,K), suggesting that CB₁R and CB₂R activation have an impact in the early stages of DG neuronal differentiation by inducing the commitment of early progenitors toward a neuronal fate.

To evaluate whether the effect upon DG neuronal differentiation involves CB₁R or CB₂R, DG cells were treated with ACEA (1 μM) or HU-308 (1 μM) or a combination of both. It was observed a significant increase in the number of NeuN-positive cells when cultures were exposed to either the CB₁R or the CB₂R selective agonists (control: 100.02 ± 0.04%; ACEA 1 μM: 154.10 ± 17.78%; HU-308 1 μM: 151.50 ± 10.61%; n = 14–16, p < 0.01), as well as when both agonists were added together (ACEA 1 μM + HU-308 1 μM: 149.90 ± 18.81%; n = 7, p < 0.05) with an effect similar to the observed with the non-selective CB₁R and CB₂R agonist WIN55,212-2 (Figures 11A,B). Furthermore, the effect caused by CB₁R selective activation was blocked either in the presence of the CB₁R selective antagonist AM251 (ACEA 1 μM + AM251 1 μM: 100.50 ± 14.17%; n = 6, p < 0.01) or in the presence of the CB₂R selective antagonist AM630 (ACEA 1 μM + AM630 1 μM: 79.42 ± 15.34%; n = 7, p < 0.001) (Figure 11C). Similarly, the effect caused by CB₂R activation was blocked either in the presence of the CB₁R selective antagonist AM251 (HU-308 1 μM + AM251 1 μM: 70.36 ± 13.58%; n = 7, p < 0.001) or in the presence of the CB₂R selective antagonist AM630 (HU-308 1 μM + AM630 1 μM: 99.34 ± 22.89%; n = 6, p < 0.01) (Figure 11D). The CB₁R selective antagonist AM251 alone was able prevent the effect of both agonists added together (ACEA + HU-308) (control: 100.04 ± 2.05%; ACEA 1 μM+HU-308 1 μM + AM251 1 μM: 72.26 ± 10.39%; n = 3, p < 0.05) (Figure 11E). Similarly, co-incubation with the CB₂R selective antagonist AM630 blocked the effect mediated by CB₁R and CB₂R co-activation (ACEA 1 μM + HU-308 1 μM + AM630 1 μM: 98.35 ± 18.60%; n = 3) (Figure 11E). Additionally, when co-incubating with both selective antagonists, the effect promoted by both selective agonists was lost, compared to control cultures (control: 100.00 ± 0.43%; ACEA 1 μM + HU-308 1 μM + AM251 1 μM + AM630 1 μM: 68.36 ± 11.52%; n = 3, p < 0.01) (Figure 11F). In any case did the incubation with the antagonists alone cause significant effects as compared with control (Figure 11). Moreover, the effect promoted by 1 μM WIN55,212-2 on DG neuronal differentiation was blocked in the presence of the CB₁R selective antagonist AM251 (WIN55,212-2 1 μM + AM251 1 μM: 74.61 ± 7.74%; n = 6, p < 0.01) (Figure 11G), as well as in the presence of the CB₂R selective antagonist AM630 (WIN55,212-2 1 μM + AM630 1 μM: 117.20 ± 15.52%; n = 7, p < 0.05) (Figure 11H). Taken together, the above results point toward an action CB₁R and CB₂R as modulators of DG neuronal differentiation and further suggest the existence of crosstalk between CB₁R and CB₂R to control postnatal neurogenesis.

To further study the cannabinoid-stimulatory effect on DG neurogenesis, we treated DG cultures with selective and non-selective CB₁R and CB₂R agonists and evaluated the effect on neurite outgrowth of βIII tubulin-positive cells (Figures 12A,B). When comparing to control cultures (Figure 12C), CB₁R and CB₂R activation with the selective and non-selective receptor agonists promoted a significant increase in the total neurite length per cell (control: 100.0 ± 0.0%; ACEA 1 μM: 154.5 ± 20.15%; HU-308 1 μM: 145.0 ± 8.45%; WIN55,212-2 1 μM: 128.7 ± 11.51%; n = 3–4; p < 0.05 and p < 0.01) (Figure 12D). Moreover, although no changes were detected regarding the number of primary neurites per neuron and the maximal neurite length (Figures 12E,F) it was observed a marked increase in the number of branch points per cell when compared to control cultures (control: 100.0 ± 0.0%; ACEA 1 μM: 141.7 ± 0.48%; HU-308 1 μM: 146.3 ± 16.70%; WIN55,212-2 1 μM: 155.7 ± 15.81%; n = 3–4; p < 0.05 and p < 0.01) (Figure 12G). These results suggest that cannabinoid receptors are important modulators of DG neuronal maturation process.

The overall crosstalk actions of CB₁R and CB₂R in SVZ and DG cultures hints at a possible physical interaction of these receptors. To test this hypothesis, we performed immunoprecipitation with an anti-CB₂R receptor antibody in SVZ and DG cultures at DIV 1 and DIV 7 (molecular weight of ∼35 and 60 kDa). Subsequent immunoblot analyses of the composition of the immunoprecipitates showed a tenuous detection of CB₁R at ∼52 and 60 kDa molecular weight in SVZ (Figure 13A) and DG (Figure 13B) cultures at both time points suggesting that CB₁R and CB₂R interact at the protein–protein physical level.