Interaction between angiotensin type 1, type 2, and mas receptors to regulate adult neurogenesis in the brain ventricular–subventricular zone

cells Article Interaction between Angiotensin Type 1, Type 2, and Mas Receptors to Regulate Adult Neurogenesis in the Brain Ventricular–Subventricular Zone Maria Garcia-Garrote 1,2, Ana Perez-Villalba 2,3, Pablo Garrido-Gil 1,2 , German Belenguer 2,4, Juan A. Parga 1,2, Francisco Perez-Sanchez 2,4, Jose Luis Labandeira-Garcia 1,2, Isabel Fariñas 2,4,* and Jannette Rodriguez-Pallares 1,2,* 1Laboratorio de Neurobiología Celular y Molecular de la Enfermedad de Parkinson, Centro Singular de Investigación en Medicina Molecular y Enfermedades Crónicas (CiMUS), Dpto. Ciencias Morfolóxicas, Universidade de Santiago de Compostela, 15782 Santiago de Compostela, Spain;maria.garcia.garr[email protected] (M.G.-G.); [email protected] (P.G.-G.); [email protected] (J.A.P.); [email protected] (J.L.L.-G.) 2Centro de Investigación Biomédica en Red de Enfermedades Neurodegenerativas (CIBERNED), 28031 Madrid, Spain; [email protected] (A.P.-V.); [email protected] (G.B.); [email protected] (F.P.-S.) 3Faculty of Psychology, Universidad Católica de Valencia, Valencia, 46100 Burjassot, Spain 4Departamento de Biología Celular, Biología Funcional y Antropología Física and Estructura de Recerca Interdisciplinar en Biotecnologia i Biomedicina (ERI BIOTECMED), Universidad de Valencia, 46100 Burjassot, Spain *Correspondence: [email protected] (I.F.); [email protected] (J.R.-P.); Tel.: +34-9635-3784 (I.F.); +34-8818-15466 (J.R.-P.) Received: 28 October 2019; Accepted: 28 November 2019; Published: 30 November 2019   Abstract: The renin–angiotensin system (RAS), and particularly its angiotensin type-2 receptors (AT2), have been classically involved in processes of cell proliferation and maturation during development. However, the potential role of RAS in adult neurogenesis in the ventricular-subventricular zone (V-SVZ) and its aging-related alterations have not been investigated. In the present study, we analyzed the role of major RAS receptors on neurogenesis in the V-SVZ of adult mice and rats. In mice, we showed that the increase in proliferation of cells in this neurogenic niche was induced by activation of AT2 receptors but depended partially on the AT2-dependent antagonism of AT1 receptor expression, which restricted proliferation. Furthermore, we observed a functional dependence of AT2 receptor actions on Mas receptors. In rats, where the levels of the AT1 relative to those of AT2 receptor are much lower, pharmacological inhibition of the AT1 receptor alone was sufficient in increasing AT2 receptor levels and proliferation in the V-SVZ. Our data revealed that interactions between RAS receptors play a major role in the regulation of V-SVZ neurogenesis, particularly in proliferation, generation of neuroblasts, and migration to the olfactory bulb, both in young and aged brains, and suggest potential beneficial effects of RAS modulators on neurogenesis. Keywords: neural stem cells; ventricular–subventricular zone; AT1 receptors; AT2 receptors; aging; proliferation; neurospheres 1. Introduction New neurons destined to integrate into olfactory bulb (OB) circuits are continuously generated from neural stem cells (NSCs) in the ventricular–subventricular zone (V-SVZ) of the lateral ventricles in the adult rodent brain. These self-renewing NSCs exhibit features that are typical of mature astroglial Cells 2019,8, 1551; doi:10.3390/cells8121551 www.mdpi.com/journal/cells Cells 2019,8, 1551 2 of 23 or fetal radial glial cells, such as expression of glial fibrillary acidic protein (GFAP), and produce transit-amplifying neural progenitor cells (NPCs). These NPCs lack self-renewal and divide a few times before they commit to differentiate into doublecortin (DCX)-positive migrating neuroblasts, which migrate anteriorly and terminally differentiate in the OB [ 1 , 2 ]. Neurogenesis is considered a complex multistage process involving proliferation, cell differentiation, and survival/integration into circuitry, finely regulated by a complex interplay between intrinsic and extrinsic factors. Neurogenesis may be broadly divided into two phases: an early phase that includes proliferation, fate commitment, and migration, and a late phase that involves synaptic development and survival of newly born neurons [ 3 ]. Additionally, neurogenesis is affected by different conditions such as stress, exercise, brain injury, or aging [ 4 , 5 ]. In fact, aging is associated with structural changes in the V-SVZ and a significant decline in neurogenesis [ 6 , 7 ]. However, the regulatory network underlying neurogenesis and the possible factors linked to its impairment with age remain poorly understood [ 6 , 8 , 9 ]. Because endogenous NSCs appear to harness the potential to repair lesioned circuits, the identification of signals and mechanisms that could stimulate NSC activity in the elderly is relevant. The renin–angiotensin system (RAS) has been classically considered a circulating humoral system with fundamental actions on blood pressure regulation and sodium and water homeostasis. The enzyme renin hydrolyses its substrate, angiotensinogen (AGT), to form the decapeptide angiotensin I (Ang I), which is biologically inactive until it is converted to angiotensin II (Ang II), the principal RAS effector, by angiotensin converting enzyme (ACE). Ang II acts through activation of two major receptors, the Ang II type-1 (AT1) and type-2 (AT2) receptors, which usually induce opposite effects [ 10 ]. In addition to the “classical” endocrine RAS, local paracrine RAS have been identified in different tissues including the brain, where all the components of a classic SRA are detected and play important functional effects [ 11 ]. AT1 receptors activate nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (NOX) and reactive oxygen species (ROS) production in different tissues including the brain [ 12 – 14 ], resulting in a pro-oxidative and pro-inflammatory environment. AT2 receptors counteract the deleterious effects of AT1 receptors (i.e., anti-oxidative, anti-inflammatory) through several mechanisms, including downregulation of AT1 expression [ 15 , 16 ]. More recently, new RAS components have been discovered, such as the anti-oxidative, anti-inflammatory Mas receptor (MasR), which adds another level of complexity to the system [17,18]. Expression of AT1 is similar before and after birth, or is even higher in the adult brain. Early descriptions have indicated AT2 receptors are very abundant in fetal tissues in agreement with actions in processes of cell proliferation, maturation, apoptosis, and regeneration [ 19 , 20 ]. Although it was classically considered that the expression of AT2 receptors decreased after birth [ 21 , 22 ], recent methodologies for their detection suggest relevant actions in the adult brain [ 23 , 24 ]. However, the potential role of RAS in adult neurogenesis, its aging-related alterations, and its possible role in regulation of neurogenesis in different neurological diseases has not been thoroughly investigated [25–29] . In the study reported here, we investigated the role of major RAS receptors on neurogenesis, particularly in proliferation, generation of neuroblasts, and migration to the OB, in the V-SVZ of both young and aged rodents. 2. Materials and Methods 2.1. Animals and In Vivo Treatments Young (3–4 months old) and aged (20–22 months old) male Sprague-Dawley rats and young (3–4 months old) and old (20 months old) male C57Bl/6J mice (wild type, WT) were used in the present study. Mice carrying genetic deletions of the Agtr1 (AT1-KO) [ 30 ] gene were obtained from The Jackson Laboratory (Bar Harbor, ME, USA) and Agt2 mutant mice (AT2-KO) [ 31 ] were generously donated by Dr. Daniel Henrion (University of Angers, Angers, France). All experiments were carried out in accordance with the Spanish (RD53/2013) and European Communities Council Directive (2010/63/EU and 86/609/CEE) and were approved by the corresponding committees at the University of Santiago Cells 2019,8, 1551 3 of 23 de Compostela and Xunta de Galicia (15005/15/002) and the University of Valencia. Rats and mice were randomly distributed in several experimental groups for the treatments. Some were treated for 15 days with the AT1 receptor antagonist candesartan cilexetil (1 mg/kg/day; AstraZeneca, Madrid, Spain) administered orally in a boluses mixed with “Nocilla” hazelnut-cream spread from Idilia Foods (Valencia, Spain) [ 32 ]. Control animals received the similar boluses without the drug. Mice treated with the AT2 receptor agonist C21 (Vicore Pharma, Gothenburg, Sweden) were intraperitoneally (i.p.) injected for 15 days at a dose of 0.3 mg/kg/day of the agonist according to previous studies [ 33 – 36 ] or an equal volume of vehicle (0.9% saline). At the last day of treatment with the drugs, rats or mice were injected i.p. with four pulses of bromodeoxyuridine (BrdU; 50 mg/kg; Sigma-Aldrich, Merck, Madrid, Spain), one every two hours, and killed 24 h after the last injection. To label BrdU-label retaining cells (BrdU-LRC), mice treated with C21 or vehicle for 15 days were injected repeatedly with seen injections of BrdU, one every 2 h, and allowed to survive for a month [37]. 2.2. Immunohistochemistry Animals were deeply anesthetized and perfused transcardially with cold 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.4. Brains were dissected out and either cryoprotected and cut with a sliding microtome or sectioned with a vibrotome into 40 µ m coronal sections. Series of free-floating sections were labelled using colorimetric single immunostaining or double immunofluorescence. For BrdU detection, sections were first denatured by incubation with 2N HCl for 15–30 min at 37 ◦ C and washed with 0.1 M sodium borate buffer (pH 8.9). Sections were preincubated with a blocking solution containing 10% normal serum in potassium phosphate-buffered saline (KPBS) with 1% bovine serum albumin (BSA; Sigma-Aldrich, San Luis, MO, USA) and 0.3% Triton X-100 (Sigma-Aldrich) for 1 h at room temperature (RT). The sections were then incubated overnight at 4 ◦ C with the corresponding primary antibodies: rabbit polyclonal antibody against AT1 receptor (1:100; sc-31181; Santa Cruz Biotechnology, Heidelberg, Germany), goat polyclonal AT2 receptor antibody (1:100; sc-9040; Santa Cruz Biotechnology), mouse monoclonal antibody (1:500; B2531; Sigma-Aldrich), or rat monoclonal antibody (1:800; ab6326; Abcam, Cambridge, UK) to BrdU; goat polyclonal antibody to DCX (1:1,000; sc-8066; Santa Cruz Biotechnology); or mouse monoclonal antibody to GFAP (1:500; MAB360; Millipore, Merck Chemicals, Darmstadt, Germany). The specificity of the AT1 and AT2 antibodies has been established in previous studies [ 38 – 40 ] and was also confirmed in our laboratory [ 23 , 41 ]. For immunoperoxidase detection, sections were washed and incubated for 1 h at RT with the corresponding biotinylated secondary antibody diluted 1:500. Sections were then incubated with avidin-biotin-peroxidase complex (ABC complex; 1:150; Thermo Fisher Scientific, Waltham, MA, USA). Finally, the labeling was visualized using 3,3 0 diaminobenzidine (DAB; Sigma-Aldrich). For immunofluorescent detection, sections were incubated with the corresponding Alexa Fluor-488, Fluor-647, or Fluor-568 secondary antibodies (Molecular Probes, Thermo Fisher Scientific, Waltham, MA, USA) for 2 h at RT. Finally, tissue sections were incubated for 30 min at RT with the DNA-binding dye Hoechst-33342 (3 × 10 −5 M in KPBS) or 4 0 ,6-diamidine-2 0 -phenylindole dihydrochloride (DAPI; 1 µ g/mL; Sigma-Aldrich), mounted on gelatin-coated slides and coverslipped with Immu-Mount (Thermo Fisher Scientific). Tissue sections were visualized with a Leica AOBS-SP5X (Leica Biosystems, Wetzlar, Germany) or an Olympus FV-10i (Olympus, Barcelona, Spain) confocal laser-scanning microscope. 2.3. Cell Counting Quantification of BrdU-immunoreactive (-ir) or DCX-ir cells was carried out in series of five sections to cover the rat V-SVZ (between Bregma coordinates 1.70 and 0.20 mm), or in series of three sections to cover the mouse V-SVZ (between Bregma coordinates 1.54 and 0.14 mm) [ 42 ]. Sampling was carried out with the computer assisted stereological toolbox (CASTGrid system; Computer Assisted Stereological Toolbox; CAST version 2.1.5.9; Olympus, Ballerup, Denmark). The V-SVZ was delineated with a 4 × magnification objective. A counting frame of 450 µ m 2 was used for cell sampling, and cell profiles were observed and counted with a 100 × oil objective (NA 1.4). The total numbers of Cells 2019,8, 1551 4 of 23 BrdU-positive or DCX-positive cells were calculated according to the optical fractionator formula [ 43 ]. The V-SVZ volume was estimated according to Cavalieri’s method [44]. 2.4. Flow Cytometry After dissection, both V-SVZs from each mouse were minced and enzymatically digested using the Neural Tissue Dissociation kit (T) (#130-093-231; Miltenyi, Bergisch Gladbach, Germany) following the instructions of the manufacturer in a gentleMACS Octo Dissociator with heaters (Miltenyi). Following addition of a trypsin inhibitor (T6522; Sigma-Aldrich) at 100 µ g/mL in 3 mL of washing medium (0.6% glucose, 0.1% NaHCO 3 , 5 mM HEPES, 2 mM l-glutamine, 0.4% BSA, 1X antibiotic/antimicotic in DMEM/F-12) and mechanical dissociation by pipetting up and down 20 times through a plastic Pasteur pipette, cell suspension was filtered (40 µ m nylon filter), pelleted (300g, 10 min), and incubated with the Dead Cell Removal kit (#130-090-101; Miltenyi) following the instructions of the manufacturer. The eluted living fraction was pelleted (300g, 10 min) and incubated in 100 µ L of blocking buffer (HBSS without calcium and magnesium, 10 mM HEPES, 2 mM EDTA, 0.1% glucose, 0.5% BSA) containing the specific primary antibodies for 30 min at 4 ◦ C: CD45-BUV395 (1:200; BD 565967; Becton Dickinson, Madrid, Spain), O4-Biotin (1:30; #130-095-895; Miltenyi), CD31-BUV395 (1:100; BD 740239; Becton Dickinson), Ter119-BUV395 (1:200; BD 563827; Becton Dickinson), Streptavidin-Alexa350 (1:200; s11249; Molecular Probes, Thermo Fisher Scientific), EGF-Alexa488 (1:300; E13345; Molecular Probes, Thermo Fisher Scientific), CD24-PerCP-Cy5.5 (1:300; BD 562360; Becton Dickinson), GLAST-PE (1:20; Miltenyi), and CD9-Vio770 (1:20, #130-102-384; Miltenyi). After washing with 1 mL of blocking buffer (300g, 10 min at 4 ◦ C), labelled samples were analyzed using an LSR-Fortessa cytometer (Becton Dickinson) with 350, 488, 561, and 640 nm lasers. Single cells were gated initially by size (FSC) and complexity (SSC) and then by SSC-A and SSC-H discordance. Prior to NSC and NPC gatings, dead cells and some non-relevant cells were excluded by DAPI (0.1 µ g/mL, added just before the analysis; D9542; Sigma-Aldrich), and staining of non-neurogenic cells using well-established markers (CD45, CD31, Ter119, O4) [45,46]. 2.5. Laser Capture Microdissection (LCM) Sections containing the mouse or rat V-SVZ were stained with a shortened protocol for neutral red (1% aqueous solution; 2 min at 4 ◦ C). LCM was performed using a PALM MicroBeam system (Zeiss, Oberkochen, Germany) controlled by PALM Robo software [ 47 , 48 ]. V-SVZ cells (or striatal cells, used as a control) were visualized under bright-field microscopy at 40 × magnification. Cell pools were selected and then cut and catapulted by laser pulses into an adhesive microtube cap (Zeiss). Cell pools were immediately lysed in RTL lysis buffer (Qiagen, Hilden, Germany) containing β -mercaptoethanol for 30 min at RT and stored at −80 ◦C. 2.6. Neurosphere Cultures NSCs were isolated from the V-SVZ of the lateral ventricle of adult young (3–4 months old), aged (20 months old) male mice, or young AT1-KO and AT2-KO mice as previously described [ 49 , 50 ]. Primary neurospheres were counted after 7 days in vitro . For passages, neurospheres were mechanically dissociated to a single-cell suspension and reseeded in growth media supplemented with the specific treatments: Ang II (100 nM; Sigma-Aldrich); ZD7155 or candesartan (AT1 receptor blockers; 1 µ M; Tocris, Bristol, United Kingdom); PD123319 (AT2 receptor blocker; 1 µ M; Sigma-Aldrich); CGP42112A (Sigma-Aldrich) or C21 (AT2 receptor agonists; 1 µ M); Ang1-7 (1 µ M; Sigma-Aldrich); and A779 (MasR blocker; 1 µ M; Bachem, Bubendorf, Switzerland). The doses were selected according to suggestions from previous studies [ 33 , 35 , 51 – 55 ]. The diameters of primary and secondary neurospheres were measured after 7 or 5 days, respectively. Cells 2019,8, 1551 5 of 23 2.7. RNA Extraction and RT-PCR In microcaptured mouse and rat cells, total ribonucleic acid (RNA) extraction was performed using the RNeasy Micro kit (Qiagen), and in neurosphere cells was performed using TRIzol (Invitrogen, Thermo Fisher Scientific) according to the manufacturer’s protocol. RNA was solubilized in 14 µ l of RNAse-free water and transcribed into complementary DNA (cDNA) using the Moloney murine leukemia virus (M-MLV; 200U; Invitrogen, Thermo Fisher Scientific) reverse transcriptase (RT). For polymerase chain reaction (PCR), a reaction mix containing the BioTaq DNA polymerase (Bioline, London, United Kingdom), NH 4 reaction buffer, MgCl 2 , dNTPs (Invitrogen, Thermo Fisher Scientific), and the specific primer pairs (Sigma-Aldrich; see Table 1) were used. Primers were designed for each gene using NCBI Primer-BLAST. PCR was performed using a MultiGene Thermal Cycler (Labnet, Madrid, Spain). Finally, the PCR products were separated using agarose gel electrophoresis and were visualized with a UV detection system (Chemidoc XRS System, BioRad, Hercules, CA, USA). The mRNA expression of AT1 and AT2 receptors was determined by real-time quantitative RT-PCR as described in previous studies [ 56 ]. Briefly, PCR was performed using a real-time QuantStudio3 platform (Applied Biosystems, Foster City, CA, USA), the EvaGreen qPCR MasterMix (Applied Biological Materials Inc., Vancouver, Canada), and the primer sequences indicated in Table 1. The data were evaluated by the delta Ct method (2 -∆∆Ct ), where Ct is the cycle threshold. Expression of each gene was obtained as relative to the housekeeping ( β -actin) transcripts, and the relative amount of mRNA was presented in the form of fold change over control. Table 1. Primer sequences used for reverse transcription polymerase chain reaction (RT-PCR). Primer Accession Number Forward Sequence (50–30) Reverse Sequence (30–50) Rat AT1R NM_030985.4 GTTAAGGGCCATTTTGTTTTTCTGG TTGTCTGGATAAATCACACAACCC AT2R NM_012494.3 AACATCTGCTGAAGACCAATAG AGAAGGTCAGAACATGGAAGG β-act NM_031144.3 TCGTGCGTGACATTAAAGAG TGCCACAGGATTCCATACC Mouse Angiotensinogen (ANG) NM_007428.4 CTGCTGGCTGAGGACAAG CGAGGAGGATGCTATTGAGAA AT1R NM_177322.3 CTCTGCTGCTCTCCCGGACTTAA AGGGCCATTTTGCTTTTCTGGGT AT2R NM_007429.5 CTGGCAAGCATCTTATGTAGTTC CAAGCATTCACACCTAAGTATTCA β-act NM_007393.5 TCGTGCGTGACATTAAAGAG TGCCACAGGATTCCATACC 2.8. Combined HPLC and Western Blot (WB) AGT was purified and detected in neurosphere growth culture media as previously described [ 57 ] with slight modifications. Culture media (1 mL) were collected, heated at 37 ◦ C, and injected (20 µ L/injection) into a Shimadzu HPLC system (Shimadzu, Kyoto, Japan). AGT was separated on a reverse phase column (Perkin Elmer Aquapore ODS 300, Perkin Elmer, Madrid, Spain) at 37 ◦ C and a flow rate of 1.5 mL/min by using a two-step separation method consisting in an initial isocratic elution (5% of acetonitrile with 0.1% phosphoric acid for 5min) and then a linear gradient from 5% to 66.5% of acetonitrile with 0.1% phosphoric acid (30 min). The AGT fraction was visualized at 240 nm with a UV-VIS detector (Shimadzu SPD-20 AV, Shimadzu), collected in a fraction collector (Shimadzu FRC-1 0 A, Shimadzu), and dried in a vacuum concentrator (Savant ISS110, Thermo Fisher Scientific). The pellet was resuspended in RIPA buffer containing a protease inhibitor cocktail (Sigma-Aldrich) and the presence of AGT in the fraction was confirmed by WB analysis. Briefly, proteins were separated using 5%–10% Bis-Tris polyacrylamide gels and transferred to nitrocellulose membranes. The membranes were incubated overnight with a goat primary antibody against AGT (1:200; sc-7419; Santa Cruz Biotechnology) and then for 1 h with the HRP-conjugated secondary antibody donkey anti-goat IgG (1:2500; sc-2020; Santa Cruz Biotechnology). Immunoreactivity was detected with a Lumminata Crescendo HRP Chemiluminescent Kit (Millipore) and visualized with a chemiluminescence detection system (ChemiDoc XRS System, BioRad, Hercules, CA, USA). AT1 and AT2 protein levels were detected in mouse V-SVZ and striatum (i.e., positive control) by WB analysis (see above) using the corresponding primary antibodies from Santa Cruz Laboratories (1:200): Cells 2019,8, 1551 6 of 23 goat polyclonal anti-AT1 (sc-31181) and rabbit polyclonal anti-AT2 (sc-9040), previously validated [ 41 ]. GAPDH (rabbit polyclonal antibody; 1:50000; G9545; Sigma-Aldrich) was used as a loading control. 2.9. Statistical Analysis Two-group comparisons were carried out by using Student’s t-test. Multiple comparisons were analyzed by one-way analysis of variance (ANOVA) followed by Bonferroni post hoc test. The normality of populations and homogeneity of variances were tested before each analysis of variance. All culture data were obtained from at least three separated experiments and normalized to the values of the control group of the same batch (100%). The results are presented as means ± standard error of the means (SEM). Differences were considered statistically significant at p<0.05. All statistical analyses were performed with SigmaPlot 11.0 (Systat Software Inc., San Jose, CA, USA). 3. Results 3.1. AT2 Receptors Mediated Promoting Effects of Ang II on Neurosphere Formation As a first approach to the analysis of local RAS in the V-SVZ murine niche, we isolated subventricular cells from young adult mice and plated them as single cells in serum-free growth medium containing mitogens EGF and FGF-2 to produce neurospheres, floating clonal aggregates that can be expanded through subculture (Figure 1A). Neurospheres can be formed by NSCs and NPCs and constitute an ideal system to evaluate the action of signaling pathways in proliferation and self-renewal [ 49 ]. The endogenous expression of AGT, AT1, and AT2 receptors by neurosphere cells was confirmed in cell lysates by PCR (Figure 1B). Moreover, the presence of the angiotensinogen (ANG) protein in the medium conditioned by the cells during 5 days was demonstrated by combining HPLC-based extraction and WB [ 57 ] (Figure 1C). Because the results indicated the possibility of RAS actions in neurospheres, we next seeded single cells dissociated from neurospheres in medium with and without 100 nM Ang II combined with concurrent administration of AT1 and AT2 receptor antagonists, 1 µ M. In particular, we used peptide antagonist ZD7155 (ZD from now on) and non-peptide antagonist candesartan to block AT1 receptors and peptide PD123319 (PD from now on) to block AT2 receptors. AT1 receptor inhibition did not produce any change in the numbers of neurospheres, but we found reduced numbers of neurospheres in the cultures treated with the AT2 receptor inhibitor, independently of the exogenous addition of Ang II, indicating that the AT2 receptor mediated promoting effects of endogenous and exogenously added Ang II in neurosphere formation (Figure 1D). We did not find, however, any significant differences in sphere diameters among treatments, suggesting that AT2 plays a role in neurosphere initiation but not growth (Figure 1E). To directly test that AT2 receptors indeed promote neurosphere formation, we plated individual neurosphere cells in the presence of two different specific agonists. Treatment with agonist CGP42112A (CGP from now on) induced an increase in the number of spheres generated, which was blocked by the AT2 receptor antagonist PD, and similar results were obtained with the non-peptide AT2 receptor agonist C21 (Figure 1F). To analyze whether the AT2 receptor-mediated positive effects on neurosphere numbers included effects on self-renewal, we re-plated cells obtained from neurospheres that had been grown in the presence of the AT2 receptor agonists and antagonists in growth medium without any specific treatment and evaluated neurosphere formation. In the cultures in which we had withdrawn the AT2 receptor modulators (i.e., derived from neurospheres pre-treated with AT2 agonist; ‘pre-treated’ condition), we could obtain information about the type of cell division that was promoted by the drugs when they were present (‘treated’ condition; Figure 1F) [ 49 ]. We observed reduced proportions of new neurospheres formed by cells that had been expanded in their previous passage in the presence of AT2 receptor agonists (Figure 1G). These data suggested that AT2 receptor activation induces asymmetrical division of neurosphere-forming cells, an effect that results in increased numbers of neurospheres that are, however, less clonogenic (i.e., asymmetrical divisions lead to a loss of stemness capacity and reduced ability to generate new neurospheres, i.e., tertiary neurospheres). Cells 2019,8, 1551 7 of 23 Cells 2019, 8, x 7 of 24 that AT2 receptor activation induces asymmetrical division of neurosphere-forming cells, an effect that results in increased numbers of neurospheres that are, however, less clonogenic (i.e., asymmetrical divisions lead to a loss of stemness capacity and reduced ability to generate new neurospheres, i.e., tertiary neurospheres). Figure 1. Generation of neurospheres from the ventricular-subventricular zone (V-SVZ) of young mice. (A) Photomicrographs showing floating neurospheres obtained from wild-type mice. (B) Representative bands for angiotensinogen (ANG), Ang II type-1 (AT1) and type-2 (AT2) receptors, and β-actin obtained by RT-PCR in neurospheres (NF). Homogenates of striatum (ST) were used as a positive control. (C) ANG was detected in neurosphere culture medium by HPLC and visualized by western blot. ANG (250 μg/mL) was used as a positive control (C+). Bar graphs showing the number (D) and diameter (E) of neurospheres after treatment with angiotensin II (Ang II), AT1 receptor antagonist (ZD7155 or candesartan), and AT2 receptor antagonist (PD123319). Histograms showing the number of neurospheres after treatment with AT2 receptor agonists (CGP42112A or C21) and the AT2 antagonist PD123319 (F; treatment) or in cultures derived from neurospheres pre-treated with AT2 agonists and reseeded in the absence of any treatment (G; neurospheres derived from cultures Figure 1. Generation of neurospheres from the ventricular-subventricular zone (V-SVZ) of young mice. ( A ) Photomicrographs showing floating neurospheres obtained from wild-type mice. ( B ) Representative bands for angiotensinogen (ANG), Ang II type-1 (AT1) and type-2 (AT2) receptors, and β -actin obtained by RT-PCR in neurospheres (NF). Homogenates of striatum (ST) were used as a positive control. ( C ) ANG was detected in neurosphere culture medium by HPLC and visualized by western blot. ANG (250 µ g/mL) was used as a positive control (C+). Bar graphs showing the number ( D ) and diameter ( E ) of neurospheres after treatment with angiotensin II (Ang II), AT1 receptor antagonist (ZD7155 or candesartan), and AT2 receptor antagonist (PD123319). Histograms showing the number of neurospheres after treatment with AT2 receptor agonists (CGP42112A or C21) and the AT2 antagonist PD123319 (( F ); treatment) or in cultures derived from neurospheres pre-treated with AT2 agonists and reseeded in the absence of any treatment (( G ); neurospheres derived from cultures pre-treated with AT2 agonists; pre- =previously treated with). All culture data were obtained from at least three separated experiments. Data are means ± standard error of the mean (SEM). * p<0.05 relative to control (untreated) group (one-way ANOVA and Bonferroni post hoc test.). SEM =standard error of the mean. ANOVA =analysis of variance. bp =base pairs. Scale bar: 150 µm. Cells 2019,8, 1551 8 of 23 3.2. AT2 Receptor Agonist C21 Stimulated Activity in the V-SVZ Niche Our data indicated that AT2 receptor activation induces proliferation of neurosphere-forming cells in the V-SVZ. We, therefore, decided to analyze the in vivo effects of the pharmacological activation of the AT2 receptor using its non-peptide agonist C21. We analyzed the cell composition of V-SVZs dissected from WT mice treated with C21 using combinations of multiple markers and FACS technology [ 46 , 58 – 60 ]. After discarding non-neurogenic cells, the remaining cell pool was sub-divided using a panel of markers that included antibodies to GLAST (astroglial cells), CD24 (neuroblasts), and CD9 (higher in NSCs than in astrocytes), as well as fluorescently-labeled EGF, which labels activated cells within the lineage, as previously described [ 45 ]. We classified the cells into EGFR + proliferating and EGFR— postmitotic GLAST—CD24 high neuroblasts, EGFR + proliferating NPCs (GLAST—CD24 low and GLAST + CD24 high cells), and CD9 high GLAST + CD24 low/neg NSCs [ 45 ] (Figure 2A,B). After a 15 day treatment with C21, we found a significant increase in the total population of NSCs and of NPCs (Figure 2C). In line with these data, we observed higher proportions of neuroblasts arriving at the OB (Figure 2C). The C21 effect was transient, as we could not observe any signs of increased activation one month after the treatment (Figure 2D). The data together further indicated that pharmacological activation of the AT2 receptor increased V-SVZ activity, and that the effect was present during a chronic treatment (i.e., 15 days) but did not continue for a long period if the treatment was stopped. Cells 2019, 8, x 8 of 24 pre-treated with AT2 agonists; pre- = previously treated with). All culture data were obtained from at least three separated experiments. Data are means ± standard error of the mean (SEM). * p < 0.05 relative to control (untreated) group (one-way ANOVA and Bonferroni post hoc test.). SEM = standard error of the mean. ANOVA = analysis of variance. bp = base pairs. Scale bar: 150 μm. 3.2. AT2 Receptor Agonist C21 Stimulated Activity in the V-SVZ Niche Our data indicated that AT2 receptor activation induces proliferation of neurosphere-forming cells in the V-SVZ. We, therefore, decided to analyze the in vivo effects of the pharmacological activation of the AT2 receptor using its non-peptide agonist C21. We analyzed the cell composition of V-SVZs dissected from WT mice treated with C21 using combinations of multiple markers and FACS technology [46,58–60]. After discarding non-neurogenic cells, the remaining cell pool was subdivided using a panel of markers that included antibodies to GLAST (astroglial cells), CD24 (neuroblasts), and CD9 (higher in NSCs than in astrocytes), as well as fluorescently-labeled EGF, which labels activated cells within the lineage, as previously described [45]. We classified the cells into EGFR+ proliferating and EGFR— postmitotic GLAST—CD24high neuroblasts, EGFR+ proliferating NPCs (GLAST—CD24low and GLAST+CD24high cells), and CD9highGLAST+CD24low/neg NSCs [45] (Figure 2A,B). After a 15 day treatment with C21, we found a significant increase in the total population of NSCs and of NPCs (Figure 2C). In line with these data, we observed higher proportions of neuroblasts arriving at the OB (Figure 2C). The C21 effect was transient, as we could not observe any signs of increased activation one month after the treatment (Figure 2D). The data together further indicated that pharmacological activation of the AT2 receptor increased V-SVZ activity, and that the effect was present during a chronic treatment (i.e., 15 days) but did not continue for a long period if the treatment was stopped. Figure 2. (A) FACS analysis of neural stem cell (NSC), neural progenitor (NP), and neuroblast (NB) populations in the ventricular–subventricular zone (V-SVZ) within the Linpopulation discriminated by levels of CD24 and GLAST. (B) FACS analysis of NSCs within the Lin-/CD24-/low cells discriminated by levels of GLAST and CD9. (C) Histogram showing the quantification of NSCs, NP, and NB FACS analyses 2 h after the last dose of 15 days C21 treatment; the levels of these three Figure 2. ( A ) FACS analysis of neural stem cell (NSC), neural progenitor (NP), and neuroblast (NB) populations in the ventricular–subventricular zone (V-SVZ) within the Linpopulation discriminated by levels of CD24 and GLAST. ( B ) FACS analysis of NSCs within the Lin-/CD24-/low cells discriminated by levels of GLAST and CD9. ( C ) Histogram showing the quantification of NSCs, NP, and NB FACS analyses 2 h after the last dose of 15 days C21 treatment; the levels of these three populations of cells raised up after 15 days of treatment. ( D ) Histogram showing FACS analyses quantification of NSC, NP, and NB populations 1 month after C21 treatment showing no differences between experimental groups. Data are means ±standard error of the mean (SEM). * p<0.05 (Student’s t-test). Cells 2019,8, 1551 9 of 23 3.3. AT1 Receptor-Dependent Restraining Action on Adult Neurogenesis Was Counteracted by Stimulatory AT2 Receptor Activity Although AT2 receptors appeared to play a significant role in the activation of neurosphere-initiating cells by endogenous Ang II, complex interactions between AT1 and AT2 receptors were observed in different tissues and, therefore, we turned to mice that carry specific deletions in the genes coding for AT1 and AT2 receptors to dissect individual and combined functions in vivo . AT1-KO and AT2-KO homozygous mutant mice and WT littermates were injected i.p. with BrdU (four pulses at 50 mg/kg, every 2 h) and sacrificed 24 h after the last injection to detect proliferating cells. Consistent with our in vitro data, we did not observe any significant change in the number of BrdU-ir and DCX-ir cells in AT1-KO mice, whereas both parameters were significantly reduced in AT2-KO mice relative to control WT mice (Figure 3A–D). These data further indicated that the AT2 receptor is a NSC positive regulator and demonstrated its essential role in vivo. Cells 2019, 8, x 10 of 24 Figure 3. Bromodeoxyuridine (BrdU (A,B) or doublecortin (DCX)-immunoreactive (-ir) (C,D) cells in the ventricular–subventricular zone (V-SVZ) of wild-type (WT) mice, AT1 receptor-deficient mice (AT1-KO), and AT2-deficient mice (AT2-KO). Number of BrdU (E) and DCX-ir (F) cells in the V-SVZ of WT mice, WT mice treated with the AT1 antagonist candesartan (Cand), WT mice treated with the AT2 agonist C21, and AT2-KO mice untreated or treated with candesartan for 15 days. Data are means ± SEM. * p < 0.05 relative to WT controls (one-way ANOVA and Bonferroni post hoc test); in figures (E) and (F), $ p < 0.05 relative to AT2-KO group (Student’s t-test). ANOVA = analysis of variance. Scale bar: 200 μm. 3.4. Cross-Regulation of AT1 and AT2 Receptors Underlies Ang II Effects on Neurosphere-Forming Cells Because our data suggested complex interactions between AT1 and AT2 receptors in the V-SVZ, and because a number of studies in different cells and tissues have shown reciprocal regulation between AT1 and AT2 receptors [61–63], we next set up to evaluate expression levels of these two receptors. We visually selected cells of the V-SVZ in lightly stained brain sections of WT, as well in AT1-KO and AT2-KO mice, which were subsequently isolated by laser-mediated capture. RT-PCR detection of AT1 and AT2 receptors with specific primers (see Table 1) in the cell pools of AT1-KO mice or of WT mice that had been treated with candesartan revealed a significant decrease in Agtr2 mRNA expression, indicating that AT1 receptor activity upregulates the expression of AT2 receptors in the V-SVZ (Figure 4A). In contrast, cells from AT2-KO mice showed a significant increase in Agtr1 mRNA expression, indicating that AT2 receptors downregulate expression of AT1 receptors in this niche (Figure 4A). In line with these results, treatment of WT neurosphere cultures with the AT2 antagonist PD increased the levels of Agtr1 mRNA, whereas the AT1 antagonist candesartan reduced Agtr2 mRNA levels (Figure 4B). Figure 3. Bromodeoxyuridine (BrdU ( A , B ) or doublecortin (DCX)-immunoreactive (-ir) ( C , D ) cells in the ventricular–subventricular zone (V-SVZ) of wild-type (WT) mice, AT1 receptor-deficient mice (AT1-KO), and AT2-deficient mice (AT2-KO). Number of BrdU ( E ) and DCX-ir ( F ) cells in the V-SVZ of WT mice, WT mice treated with the AT1 antagonist candesartan (Cand), WT mice treated with the AT2 agonist C21, and AT2-KO mice untreated or treated with candesartan for 15 days. Data are means ± SEM. * p<0.05 relative to WT controls (one-way ANOVA and Bonferroni post hoc test); in figures ( E , F ), $ p<0.05 relative to AT2-KO group (Student’s t-test). ANOVA =analysis of variance. Scale bar: 200 µm. Cells 2019,8, 1551 16 of 23 The results also show that AT1 and AT2 receptors played a major role in NSC proliferation and/or generation of neuroblasts in the V-SVZ. Most of known actions of Ang II in the adult brain were mediated by AT1 receptors [ 74 ]. Local Ang II, via AT1 receptors, has been shown to enhance oxidative stress damage and neuroinflammation contributing to the progression of neurodegeneration [ 75 , 76 ]. However, effects of AT1 on cell proliferation and neurogenesis are controversial. Ang II, through AT1 receptors, is a major activator of the NADPH-oxidase complex, which is a primary source of ROS [ 77 ], and enhanced ROS generation plays a recognized role in cell death and pathogenesis. However, previous studies have also shown that ROS derived from NADPH-oxidase activation contribute to the self-renewal [ 78 ] and lineage determination in NSCs [ 79 ], and that AT1 receptors have an important role in progenitor cell proliferation [ 80 ]. Other studies have observed that blockage of AT1 receptors with different antagonists such as candesartan, valsartan, or losartan induced proliferation and neurogenesis in the dentate gyrus [ 25 , 26 , 81 ], but also that treatment with AT1 antagonists or angiotensin converting enzyme inhibitors diminishes neurogenesis [ 27 , 28 , 82 – 84 ], or that AT1 blockade does not affect hippocampal neurogenesis [ 85 , 86 ]. There are also controversial results on the effects of AT2 receptors. Activation of AT2 receptors with the selective agonist CGP reportedly induced an increase in BrdU-positive cells in models of traumatic brain [ 87 , 88 ] and proliferation of hippocampal NSCs in vitro [ 68 ]. However, antiproliferative effects of AT2 receptors have also been suggested [ 89 – 91 ]. Our results suggest that most of the aforementioned controversial results may be explained by the complex interactions between AT1 and AT2 receptors and possible differences in the AT1/AT2 ratio in different regions and species. In AT2-KO mice, which show increased expression of AT1 in the V-SVZ, we observed a marked decrease in proliferation and generation of neuroblasts, which was inhibited by treatment with the AT1 antagonist candesartan, suggesting that a decrease in AT2 activity and an increase in AT1 activity inhibit proliferation and generation of neuroblasts. This was confirmed by treating WT mice with the AT2 agonist C21, which increased proliferation and generation of neuroblasts. Interestingly, AT1-KO mice or WT mice treated with the AT1 blocker candesartan, which also showed downregulation of AT2 receptors, did not show significant changes in proliferation and generation of neuroblasts. A basal inhibition of proliferation by AT1 receptors without any direct effect of AT2 receptors (i.e., beyond inhibition of AT1 activity) should lead to higher levels of proliferation in AT1-KO mice relative to WT mice. These similar levels may be explained by an additional direct positive effect of AT2 receptors on proliferation, which is inhibited by the downregulation of AT2 receptors observed in AT1-KO and candesartan-treated mice or neurosphere cultures. In summary, both downregulation of AT1 and upregulation of AT2 contribute to the increase in V-SVZ proliferation and generation of neuroblasts. The results obtained in cultures of neurospheres were consistent with that observed in mice. Activation of AT2 receptors led to an increase in the generation of neurospheres. On the contrary, inhibition of AT2 led to a decrease in generation of neurospheres, and no significant changes were observed after blocking AT1 (which also leads to AT2 downregulation in neurospheres) as observed in neurospheres from AT1-KO mice or candesartan or ZD-treated WT mouse neurospheres. We also observed that the generation of neurospheres was reduced after previous stimulation of AT2 receptors, suggesting a possible loss of stemness. The rat V-SVZ and mouse neurospheres from aged animals showed a significant decrease in proliferation and neurogenesis, which was inhibited by AT2 agonists or AT1 antagonists and exacerbated by AT2 antagonists. These results are consistent with the aforementioned mechanisms, as we observed an increase in AT1/AT2 expression ratio in the V-SVZ of aged animals relative to young controls, as previously observed in different types of cells [ 92 ], and particularly in the striatum and substantia nigra [ 62 , 65 , 66 ]. The present results are consistent with previous studies showing that neurogenesis continues throughout life, although its rate declines with increasing age in rodents and non-human primates [ 64 ], and with previous studies showing a progressive reduction in the production of progenitors throughout the mouse adult life and a maintenance of NSC pools [ 6 , 7 , 93 ]. Interestingly, our results reveal that the reduction in proliferation and generation of neuroblasts and Cells 2019,8, 1551 17 of 23 neuropheres observed in the V-SVZ of aged animals may be counteracted by acting on the local RAS, suggesting that old NSCs show proliferation and differentiation capacity after adequate stimulation, which is consistent with other recent observations [94]. In cultures of neurospheres, we also investigated a possible role of Ang1-7/MasR. We administered Ang 1–7 to cultures derived from young and aged mice, and no significant change was detected. However, treatment of cultures with the MasR antagonist A779 blocked the AT2 agonist-induced increase in the number of spheres. This suggests a functional dependence between AT2 and MasR for the AT2-induced generation of neurospheres. Accumulating evidence shows that both receptors are co-localized, form heterodimers, and have different functional interactions in different cells and experimental conditions [ 27 , 95 – 97 ], and recent data from our laboratory (unpublished) showed that young AT2-KO mice show a reduced expression of MasR in the substantia nigra and striatum, which may also contribute to the decrease in proliferation and neurogenesis observed in the V-SVZ of AT2-KO mice. Therefore, although RAS receptors have been associated with different cellular and physiological responses, their role in neurogenesis is complex, as shown by the present results that reveal the presence of AT1, AT2, and MasR interactions possibly both in physiological or pathological situations. 5. Conclusions The present data revealed that the RAS played a major role in the regulation of adult neurogenesis both in young and aged brains. Ang II, via AT1 receptors, set a basal inhibition of V-SVZ proliferation that was antagonized by AT2 receptors. In addition, AT2 receptor activity may exert an additional direct stimulatory effect on proliferation. AT1/AT2 expression and ratio were affected by aging, which resulted in a decrease of neurogenesis. However, AT1 and AT2 receptor expression and neurogenesis may be modulated by therapies with agonists and antagonist of these receptors. Furthermore, RAS may interact with other modulators of neurogenesis. It is well-known that dopamine modulates neurogenesis in the V-SVZ [ 98 ], and counterregulatory interactions between Ang II and dopamine receptors have been demonstrated in the striatum [ 57 , 66 ] and other tissues [ 99 , 100 ]. Therefore, it seems plausible that dopamine regulates proliferation and neurogenesis by modulating RAS activity. Our results also suggest the potential beneficial effects of RAS modulators on neurogenesis. However, AT1/AT2 expression, ratio, and responses to drugs may vary depending on the species. Future studies are required to further clarify the effects of manipulation of the brain RAS in humans and pathological conditions associated with neurogenesis impairments, and to provide a solid basis for the use of RAS manipulation in such diseases. Author Contributions: J.R.-P., J.L.L.-G., and I.F. designed the research; M.G.-G., A.P.-V., P.G.-G., J.A.P., G.B., and J.R.-P. performed the experiments. All authors contributed to the editing of the manuscript. Funding: This research was funded by Spanish grants from Ministerio de Econom í a y Competitividad (BFU2015-70523 and SAF2017-86690-R), Instituto de Salud Carlos III (Retic TERCEL RD16/0011/0016, RD16/0011/0017, and CIBERNED), Galician Government (XUGA, ED431C2018/10; ED431G/05), FEDER (Regional European Development Fund), Generalitat Valenciana (Prometeo 2017-030), and Fundaci ó n Emilio Bot í n-Banco Santander. The funding sources had no involvement in the design, data collection, analysis and interpretation, writing, or submission of the study. Acknowledgments: We thank Pilar Aldrey, Iria Novoa, and Cristina Gianzo for their technical assistance. Conflicts of Interest: The authors declare no conflict of interest. References 1. Obernier, K.; Alvarez-Buylla, A. Neural stem cells: Origin, heterogeneity and regulation in the adult mammalian brain. Development 2019,146. [CrossRef] 2. Lim, D.A.; Alvarez-Buylla, A. 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