To understand the role of Shh signaling in seizure-induced aberrant neurogenesis, we first investigated whether Shh signaling is activated upon induction of seizure activity. Expression of Gli1, a downstream transcription factor of Shh signaling, is transcriptionally induced upon activation of Shh signaling (Bai et al., 2002; Lee et al., 1997). In the subgranular zone (SGZ) of the DG, Gli1 expression and activation of Shh signaling are seen exclusively in NSCs (Ahn and Joyner, 2005; Bottes et al., 2021). To confirm this, using Gli1^CreER mice crossed with Cre-dependent red fluorescent protein (tdTomato) reporter mice Rosa^AI14 (Ahn and Joyner, 2004; Madisen et al., 2010), we labeled Gli1 expressing cells in the DG with tdTomato. One day after a 3-day tamoxifen treatment, we observed that about 90% of tdTomato-labeled cells in the SGZ were Sox2+ S100B− cells (Figure 1—figure supplement 1), comprising Sox2+ GFAP+ radial glial cell population. This confirms that NSCs are Shh responsive and that Gli1 expression in the SGZ is essentially restricted to NSCs. Thus, we used Gli1^nLacZ/+ mice, which carry the nuclear LacZ (nLacZ) transgene driven by the promoter of Gli1 (Ahn and Joyner, 2005; Bai et al., 2002; Ihrie et al., 2011) as a sensitive readout for the activation of Shh signaling in NSCs and examined the number of Gli1-nLacZ+ cells, which represent Shh-responding cells, in SGZ after seizure induction. We induced seizures with consecutive intraperitoneal injection of low-dose kainic acid (KA) in Gli1^nLacZ/+ mice and found that Gli1-nLacZ+ Shh-responding cells in the SGZ are significantly increased 3 days post-seizure induction (Figure 1A, B). These data show that Shh signaling activity is increased in NSCs after KA-induced seizures.

To investigate whether Shh contributes to seizure-induced aberrant neurogenesis, we analyzed Shh knock-in heterozygote (Shh^EGFP-Cre/+, from here on referred as Shh^+/−) mice (Harfe et al., 2004). We found that Gli1-nLacZ+ Shh-responding cells in the SGZ of Shh^+/−; Gli1^nLacZ/+ mice were significantly reduced (Figure 1C, D), indicating that the level of Shh signaling in NSCs is decreased in these mice. We also observed that Gli1-nLacZ+ cells failed to increase after seizure induction in these mice, indicating that Shh signaling activity is not upregulated with seizure activity in Shh^+/− mice. We then investigated whether seizures induced aberrant neurogenesis in Shh^+/− mice. After induction of seizures by KA, we injected 5-bromo-2′-deoxyuridine (BrdU) for 5 days to label newborn neurons and analyzed the number of BrdU-labeled DCX+ newborn neurons at 3 days after the last BrdU injection (Figure 1E). In WT mice, the number of BrdU+ DCX+ newborn neurons dramatically increased after seizure induction. However, this was significantly reduced in Shh^+/− mice by comparison (Figure 1F–H). Ectopic migration of newborn neurons into the hilus is known to occur in post-seizure neurogenesis (Lybrand et al., 2021; Parent et al., 1997). Shh^+/− mice have a reduction in ectopic hilar dentate neurons (Figure 1I). These results demonstrate that Shh contributes to seizure-induced aberrant neurogenesis.

To further understand the role of Shh in seizure-induced aberrant neurogenesis, we sought to identify the source of Shh ligands in the DG, particularly after seizures. Shh^EGFP-Cre/+ mice were crossed with Rosa^AI14 reporter mice (from here on referred as Shh^Cre/+;Rosa^AI14) and seizures were induced. One week after seizure induction, we found that there was specific tdTomato expression in the dentate hilus of Shh^Cre/+;Rosa^AI14 mice both with and without seizure induction (Figure 2A, B). Since inhibitory interneurons and mossy cells are both prominent dentate hilar cell types, we explored which of these are included in the tdTomato+ cells. Co-immunostaining with the cell-type markers showed that the majority of tdTomato+ cells were GluR2/3+ mossy cells but with a much smaller number of Parvalbumin+ interneurons expressing tdTomato (Figure 2C, D). Furthermore, the cell types reflected in the population of tdTomato+ cells did not change with seizure induction, thus indicating that in the hilus, seizures did not lead to ectopic expression of Shh in cells that normally do not express the ligand (Figure 2D). Interestingly, we found that tdTomato+ cells were dominantly observed in the dorsal GluR2/3+ mossy cells but not in the ventral hippocampus (Figure 2—figure supplement 1). These data suggest that dorsal mossy cells are the major source of Shh in DG in both the naive and post-seizure hippocampus. We also observed increased numbers of Shh-responding NSCs of the SGZ upon seizure induction (Figure 1A, B). Therefore, we next tested the possibility that the amount of Shh expression is upregulated by seizures in mossy cells. To address this, we visualized Shh mRNA in the DG using RNAscope in situ hybridization technology. Consistent with the results from Shh^Cre/+;Rosa^AI14 mice, we observed that Shh mRNA is enriched in the dentate hilus (Figure 2E). Furthermore, Shh expression was dramatically increased throughout the dentate hilus from anterior to posterior of dorsal DG at 24 hr after seizure induction. Combining RNAscope with immunofluorescence, we analyzed the number of Shh mRNA puncta in the GluR2/3+ mossy cells and found significantly increased Shh in mossy cells after seizure induction (Figure 2F, G). Taken together, these data support the conclusion that mossy cells are the local source of Shh in the DG and that Shh mRNA expression is increased upon seizure activity.

To examine the role of Shh derived from mossy cells in seizure-induced activation of Shh signaling, we generated mossy cell-selective conditional Shh knock-out (Shh-cKO) mice by crossing a floxed Shh allele (Shh^fl) with calcitonin receptor-like receptor (Crlr)-Cre mice, in which Cre-mediated recombination is induced in mossy cells and a subset of CA3 neurons in the hippocampus (Jinde et al., 2012). We first tested whether deletion of Shh in mossy cells affects Shh signaling using Gli1^nLacZ/+ mice. We found that the number of Gli1-nLacZ+ cells was significantly decreased in Shh-cKO mice compared with control mice (Crlr-Cre;Shh^+/+ mice and Shh^+/+, Shh^fl/+ and Shh^fl/fl without the Cre transgene were used as controls) (Figure 3A, B). Three days after seizure induction, Gli1-nLacZ+ cells were significantly increased in control mice. However, this increase was not observed in Shh-cKO;Gli1^nLacZ/+ mice, in which Gli1-nLacZ+ cells were rarely detected in the SGZ. We found that there were comparable number of Sox2+ GFAP+ radial NSCs in Shh-cKO mice compared with control mice (Figure 3—figure supplement 1), indicating that reduction of Gli1-nLacZ+ cells is not due to general loss of NSCs. These data suggest that Shh from mossy cells is responsible for Shh signaling in the DG, and that Shh signaling fails to be upregulated by seizures in the absence of Shh from mossy cells.

Since we found impairment in seizure-induced activation of Shh signaling in Shh-cKO mice, we next investigated whether Shh from mossy cells is involved in seizure-induced aberrant neurogenesis. The mice were treated with BrdU for 5 days, starting 1 day after seizure induction, and analyzed for the number of BrdU-labeled DCX+ newborn neurons at 3 days after the last BrdU injection (Figure 3C). We found that the number of DCX+ BrdU+ newborn neurons was comparable between control and Shh-cKO mice in the KA treatment-naive condition, suggesting that deletion of Shh from mossy cells does not significantly affect baseline neurogenesis in this timeframe. After seizure induction, the number of BrdU+ DCX+ newborn neurons was significantly increased in both groups. However, we found that the number of BrdU+ DCX+ newborn neurons was significantly reduced in Shh-cKO mice compared to control mice after seizure induction (Figure 3D, E), indicating that deletion of Shh in mossy cells attenuates seizure-induced neurogenesis. Together, these data indicate that mossy cells function as a critical source of Shh for Shh signaling activity of NSCs, and that Shh derived from mossy cells contributes to seizure-induced neurogenesis.

A previous study showed that neuronal activity triggers secretion of Shh from hippocampal neurons in culture (Su et al., 2017). Since mossy cells undergo hyperactivation during seizures, we hypothesized that once activated by seizures, mossy cells increase secretion of Shh and contribute to seizure-induced aberrant neurogenesis. To test whether neuronal activity in mossy cells increases neurogenesis, we used a chemogenetic approach to activate mossy cells. Crlr-Cre mice were crossed with Cre-dependent Designer Receptors Exclusively Activated by Designer Drugs (DREADD) activator mice (Rosa^DIO-hM3Dq) which express excitatory G-protein-coupled receptor hM3Dq with mCherry upon Cre recombination (from here on referred as Crlr-Cre;hM3Dq) (Sciolino et al., 2016; Figure 4A). In Crlr-Cre;hM3Dq mice, mCherry+ recombined cells were observed specifically in the dentate hilus throughout dorsal and ventral hippocampus (Figure 4B and Figure 4—figure supplement 1). As previously reported, we found that over 90% of mCherry+ expressing cells are GluR2/3+ mossy cells (Jinde et al., 2012), and that these made up 65% of the GluR2/3+ mossy cells in the dentate hilus (Figure 4C, D). Since the high dose of Clozapine-N-oxide, frequently used in the DREADD system has a risk of causing behavioral abnormalities and might have contributory effects on neurogenesis (Gomez et al., 2017; MacLaren et al., 2016), we instead used Clozapine (CLZ) to induce neuronal activation (Cho et al., 2020; Gomez et al., 2017). In addition, we administrated CLZ to both Rosa^DIO-hM3Dq control and Crlr-Cre;hM3Dq mice to normalize the possible side-effects of CLZ administration. 1.5 hr after CLZ injection, we observed that expression of c-fos, a marker of neuronal activation, was significantly increased in the hilus of Crlr-Cre;hM3Dq mice (Figure 4E, F); we also confirmed that over 70% of GluR2/3+ mCherry+ mossy cells were activated (Figure 4G). To investigate the effect of mossy cell neuronal activity on neurogenesis, the mice received BrdU for 5 days concomitant with CLZ treatment (Figure 4H) and were analyzed 3 days after the last BrdU injection. We found that the number of DCX+ BrdU+ newborn neurons was significantly increased in Crlr-Cre;hM3Dq mice (Figure 4I, J), suggesting that increased neuronal activity of mossy cells drove neurogenesis. We therefore next investigated whether Shh signaling activity is activated by mossy cell neuronal activity using Gli1^nLacZ/+ mice. After 6 days of CLZ administration (Figure 4K), we found that Gli1-nLacZ+ were significantly increased in the SGZ of Crlr-Cre;hM3Dq;Gli1^nLacZ/+ mice, compared with Rosa^DIO-hM3Dq;Gli1^nLacZ/+ control mice (Figure 4L, M), suggesting that Shh signaling activity is upregulated in NSCs by mossy cell neuronal activity. To test the role of Shh in mossy cell neuronal activity-induced neurogenesis, we deleted Shh in mossy cells and investigated whether neuronal activity in mossy cells induces neurogenesis using Crlr-Cre;Shh^fl/fl;hM3Dq mice. After CLZ administration, we found no induction of neurogenesis by mossy cell neuronal activity in Crlr-Cre;Shh^fl/fl;hM3Dq mice (Figure 4N, O), indicating that induction of neurogenesis by mossy cell neuronal activity is compromised in the absence of Shh from mossy cells. Together, these data indicate that Shh from mossy cells contributes to neuronal activity-induced neurogenesis by mossy cells.

Since mossy cells project their axons to the contralateral DG (Botterill et al., 2021; Houser et al., 2021), we then examined whether mossy cell neuronal activity influences neurogenesis in the contralateral DG. To address this, we stereotaxically injected Cre-dependent adeno-associated virus (AAV) expressing the excitatory G-protein-coupled receptor hM3Dq-mCherry under the human synapsin promoter (AAV::hSyn-DIO-hM3Dq-mCherry, from here on referred as AAV::hM3Dq-mCherry) into the DG of Crlr-Cre mice (Figure 4—figure supplement 2A). The mice were treated with CLZ to activate hM3Dq and analyzed 1.5 hr after CLZ injection. We found the specific infection of dorsal mossy cells in the AAV-injected side of the DG and contralateral projection of infected mossy cells in the inner and middle molecular layer in the contralateral DG (Figure 4—figure supplement 2B–D). We also confirmed that CLZ injection increases the expression of c-fos in the AAV::hM3Dq-mCherry infected mossy cells of the DG on the injected side (Figure 4—figure supplement 2E–G). To test whether mossy cell neuronal activity induces neurogenesis in contralateral DG, we injected AAV virus at 4–5 weeks old, and after 4 weeks recovery, mice received BrdU for 5 days concomitant with CLZ treatment and were analyzed 3 days after the last BrdU injection (Figure 4—figure supplement 2H). We found that the number of newborn neurons was significantly increased on the contralateral DG of AAV::hM3Dq-mCherry-injected mice (Figure 4—figure supplement 2I, J), indicating that mossy cell neuronal activity contributes to increasing neurogenesis in the contralateral DG.

Previous studies showed that Shh can be secreted from axons and anterogradely transported to distal regions along the axon (Beug et al., 2011; Peng et al., 2018; Su et al., 2017). These reports, together with our findings, raise the possibility that increased Shh expression after seizures is transported to the contralateral DG, thereby allowing mossy cells to regulate contralateral neurogenesis with neuronal activity. To address this possibility, we deleted Shh in mossy cells in a unilateral hemisphere of DG by injecting AAV::Cre-P2A-tdTomato (AAV::Cre-tdTomato) into the DG of Shh^fl/fl mice at 4–5 weeks old and investigated seizure-induced neurogenesis in the contralateral DG in 2-month-old mice (Figure 4—figure supplement 3A). We found that infection by AAV::Cre-tdTomato was restricted to the injected DG. Dorsal mossy cells send projections to the inner and middle molecular layers of DG bilaterally into both hemispheres along the septo-temporal axis (Botterill et al., 2021; Houser et al., 2021). We found tdTomato+ mossy cell projections in the contralateral DG from the DG ipsilateral to the injection site (Figure 4—figure supplement 3B), indicating that AAV::Cre-tdTomato infected mossy cells. Shh expression is observed predominantly in dorsal mossy cells (Figure 2—figure supplement 1). We observed AAV::Cre-tdTomato infection in 31% of total GluR2/3+ mossy cells and 45% of GluR2/3+ dorsal mossy cells in the DG ipsilateral to the injection site (Figure 4—figure supplement 3C, D). We then tested whether Shh deletion affects contralateral neurogenesis (Figure 4—figure supplement 3E) and found that seizure-induced neurogenesis was attenuated in the contralateral DG of mice injected with AAV::Cre-tdTomato, in which DCX+ BrdU+ newborn neurons induced by seizure activity were significantly reduced compared with mice injected with AAV::mCherry control virus (Figure 4—figure supplement 3F, G). This suggests that mossy cells control contralateral neurogenesis by regulating Shh expression after seizures. Together with our findings that Shh expression was increased by seizure (Figure 2F, G), these data demonstrate that increased Shh following seizures may be transported and contribute to neurogenesis in the contralateral DG. Taken together, these data support our hypothesis that mossy cells release Shh with increased neuronal activity and induce neurogenesis driven by seizures (Figure 4P).

Whether there is a functional benefit of increased Shh expression in mossy cells following seizure activity remains unclear, considering that seizure-induced aberrant neurogenesis negatively affects hippocampal function. Recently, it was shown that a population of quiescent NSCs in the adult DG, once activated to proliferate and produce neurons, can return to quiescence rather than being consumed by terminal differentiation (Botterill et al., 2021; Harris et al., 2021). This mechanism is suggested to contribute to the long-term persistence of NSCs and maintenance of neurogenesis throughout life by decelerating NSC pool exhaustion. Since Shh is important for proliferation and self-renewal of NSCs (Ahn and Joyner, 2005; Choe and Pleasure, 2013; Lai et al., 2003), we investigated whether Shh from mossy cells is involved in maintenance of NSCs after proliferation. We labeled proliferating NSCs with BrdU in the drinking water for 5 days after seizure induction (Figure 5A). Four weeks after BrdU administration, we analyzed the number of NSCs retaining BrdU, as previously addressed (Harris et al., 2021). Proliferated NSCs labeled by BrdU are eliminated from the NSC pool once they differentiate into neurons, whereas NSCs that have returned to the stem cell state after proliferation will be detected as BrdU-retaining Sox2+ GFAP+ radial NSCs even 4 weeks after proliferation (Figure 5B). We found that seizure induction increases the number of BrdU+ Sox2+ proliferating NSCs at day 0 (last day of BrdU labeling) in both control and Shh-cKO mice compared with the no seizure condition, although it was significantly attenuated in Shh-cKO mice compared with control mice (Figure 5C, D). Four weeks after BrdU labeling, we found that the number of BrdU-retaining Sox2+ GFAP+ radial NSCs was increased in seizure-induced mice compared with no seizure-induced mice, as expected from the increased proliferation after seizure (Figure 5C, E). However, the number of BrdU-retaining Sox2+ GFAP+ radial NSCs in Shh-cKO mice was significantly reduced compared with seizure-induced control mice (Figure 5C, E). To assess the persistence of NSCs’ identity after seizure-induced proliferation, we calculated the fraction of NSCs that persist as NSCs 4 weeks after proliferation by dividing the number of BrdU-retaining Sox2+ GFAP+ radial NSCs at 4 weeks with the number of BrdU+ Sox2+ proliferating NSCs at day 0 (Figure 5F). We found that the fraction of NSCs that retain stem cell identity after proliferation was comparable in control mice with and without seizure induction. This indicates that even though seizures induce NSCs to proliferate, they return to the stem cell state in the same ratio (Figure 5F). In contrast, we found that the fraction of NSCs that returned to the stem cell state in Shh-cKO mice was significantly reduced after seizure-induced proliferation (Figure 5F), suggesting that NSCs are less likely to return to their stem cell state after seizure-induced proliferation in the absence of Shh from mossy cells.

Seizures increase NSC proliferation and subsequent neurogenesis in the short term. However, it has been suggested that increased neurogenesis induced by neuronal hyperactivity accelerates consumption of NSCs and leads to depletion of NSC pool over time (Fu et al., 2019; Sierra et al., 2015). Since NSCs in Shh-cKO mice had fewer NSCs that returned to their stem cell state after seizure-induced proliferation, we next investigated the consequences on the NSC pool after seizure-induced neurogenesis. We analyzed the number of Sox2+ GFAP+ radial NSCs 4 weeks after BrdU labeling (which is 5 weeks after seizure induction). We found that the number of Sox2+ GFAP+ radial NSCs in the SGZ of control mice is comparable with and without seizures, suggesting that the NSC pool size is maintained at least 5 weeks after seizure induction in control mice (Figure 5G, H). However, in Shh-cKO mice, the number of Sox2+ GFAP+ radial NSCs was significantly reduced 5 weeks after seizure induction (Figure 5G, H), suggesting that deletion of Shh in mossy cells leads to premature depletion of the NSC pool after seizure-induced neurogenesis. Taken together, these data imply that Shh from mossy cells is required for maintaining the NSC pool after seizure-induced neurogenesis, and that Shh from mossy cells contribute to the NSCs’ return to their stem cell state after seizure-induced proliferation, thus preventing NSC pool depletion.

NSCs in the adult DG self-renew and return to a quiescent state after proliferation more often in aged mice than in young mice, which is thought to be the mechanism for maintaining the NSC pool during the aging process (Harris et al., 2021). We therefore next investigated whether deletion of Shh in mossy cells influences the aging-related NSC pool decline using 9- to 11-month-old aged mice. We found that the number of Sox2+ GFAP+ radial NSCs was significantly reduced in Shh-cKO mice compared with control mice (Figure 6A, B). We next asked whether the aging-related increase in NSC self-renewal is compromised in Shh-cKO mice. We administrated BrdU to mice via drinking water for 5 days to label proliferating cells and investigated the number of NSCs retaining BrdU in the SGZ at 10 days and 4 weeks after BrdU administration (Figure 6C). Consistent with previous reports (Harris et al., 2021), we found that a greater fraction of NSCs retained BrdU until 4 weeks after BrdU labeling in aged control mice compared with 2-month-old mice (Figures 5C, F, 6D, E), indicating that NSCs in aged mice tend to retain their stem cell state after proliferation compared to younger mice. However, in Shh-cKO mice, the fraction of NSCs retaining BrdU after BrdU labeling was significantly reduced compared with control mice (Figure 6D, E), suggesting that NSCs in aged Shh-cKO mice are less able to retain their stem cell state after proliferation. Together, our data suggest that Shh from mossy cells contributes to persistence of the NSC state after proliferation throughout life, which sustains the NSC pool during aging.

All mice used in this study were maintained on a 12-hr light/dark cycle with free access to food and water. Mouse colonies were maintained at University of California San Francisco (UCSF) in accordance with National Institutes of Health and UCSF guidelines. The following mouse lines were obtained from Jackson Laboratory (Bar Harbor, Maine): Gli1^CreERT2/+ (stock #007913, RRID:MGI:3053957), Rosa^AI14 (Stock #007908, RRID:MGI:J:155793), Gli1^nLacZ/+ (stock #008211, RRID:MGI:J:79392), Shh^EGFP-Cre/+ (stock #005622, RRID:MGI:3053959), Shh^flox/flox (stock #004293, RRID:MGI:1934268), and Crlr-Cre (stock #023014, RRID:MGI: 5523525). Rosa-CAG-LoxP2272-GFP-LoxP-mCherry-hM3Dq-LoxP2272-LoxP (Rosa^DIO-hM3Dq) mice were generously provided by Dr. Zachary Knight (University of California, San Francisco, also available from Jackson laboratory, stock # 026943, RRID:MGI: 5771785). All mice were randomly assigned to experiments. Both male and female mice were analyzed with no distinction, except for seizure induction experiments, where only male mice were used. There was no difference in the number of newborn neurons and radial NSCs between Crlr-Cre;Shh^fl/fl and Crlr-Cre;Shh^fl/fl;Rosa^DIO-hM3Dq at 2 and 9–11 months old without clozapine administration (Figure 6—figure supplement 1). Thus, in the study of long-term tracing of NSCs in aged mice, both Crlr-Cre;Shh^flox/flox and Crlr-Cre;Shh^flox/flox;Rosa^DIO-hM3Dq mice were used as Shh-cKO mice.

One-month-old mice were anesthetized with Isoflurane (Piramal) in the sealed box and placed on stereotaxic instrument (Kopf). Anesthesia was maintained during surgery using a veterinary vaporizer (Surgivet). Unilateral stereotaxic injections were performed into the right dorsal dentate hilus (anteroposterior: −2.0 mm from Bregma, mediolateral: 1.9 mm, dorsoventral: 2.25 mm). All injections were performed using a Nanojector III (Drummond Scientific) with a glass capillary, pre-pulled by vertical micropipette puller (Sutter Instrument company, P-30). 500 nl of virus was injected at a rate of 60 nl/min. After the virus was delivered, the glass capillary was left in place for another 5 min to allow for diffusion of the virus away from the needle tract and was then slowly withdrawn. The mice recovered in a warm cage after suturing of the incision and then returned to their home cages. After 3–4 weeks of recovery, mice were subjected to the experiments at 2 months old. For Shh deletion experiments, either AAV-hSyn-mCherry (Addgene, 114472-AAV5) or AAV-hSyn-Cre-P2A-dTomato (Addgene, 107738-AAV5) were injected at a titer of 3.0 × 10¹¹ GC/ml. For chemogenetic activation of mossy cell experiments, either AAV-hSyn-DIO-mCherry (Addgene, 50459-AAV8) or AAV-hSyn-DIO-hM3D(Gq)-mCherry (Addgene, 44361-AAV8) were injected at a titer of 4.0 × 10¹² GC/ml.

Seizure induction was performed by consecutive intraperitoneal injection of low-dose KA (Enzo Life Sciences, dissolved in saline (0.9% NaCl)) in 2-month-old male mice. Mice received KA initially at a dose of 5 mg/kg. Then 20 min after initial injection, the mice consecutively received 2.5 mg/kg KA injection every 20 min until an initial Racine stage 4/5 seizure was observed, which are defined as rearing with forelimb clonus (stage 4); rearing and falling with forelimb clonus (generalized motor convulsions, stage 5) (Racine, 1972). Mice that did not display stage 4/5 seizure after receiving total 30 mg/kg of KA were excluded from the experiments. Seizure induction and behavior measurement were performed blindly to the genotype. Mice receiving no KA treatment were used as naive mice controls.

Tamoxifen (Sigma) was dissolved in corn oil with 10% of ethanol at 50 mg/ml. Gli1^CreER/+;Rosa^AI14 mice were orally administered 5 mg of tamoxifen once a day for 3 days at 2 months old and analyzed 1 day after last tamoxifen injection. For analyzing neurogenesis by BrdU pulse labeling, mice were intraperitoneally injected with BrdU (Sigma) dissolved in saline at a dose of 50 mg/kg once a day for 5 days at 2 months old and analyzed 3 days after last BrdU injection. For long-term tracing of BrdU-labeled NSCs in the experiments for NSC persistence analysis, mice were provided 1 mg/ml of BrdU in the drinking water for 5 days at 2 or 9- to 10 months old and analyzed on the last day of BrdU administration (day 0), or 10 days or 4 weeks after BrdU administration. For induction of neuronal activity by DREADD-based chemogenetic tools, mice were administrated with clozapine (CLZ, Sigma) by intraperitoneal injection at a dose of 0.1 mg/kg once a day and by drinking water (0.1 mg/100 ml) for 6 days. CLZ drinking water was replaced every 2 days. In the experiments testing induction of neuronal activity by c-fos expression, mice were perfused at 1.5 hr after intraperitoneal injection of 0.1 mg/kg of CLZ. For the analysis of Shh-responding cells using Gli1^nLacZ/+ mice in DREADD experiments, mice were given CLZ for 6 days and perfused on the last day of CLZ administration. To investigate neurogenesis in DREADD experiments, mice were given CLZ for 6 days, and starting on the second day of CLZ administration, mice were also given 50 mg/kg of BrdU by intraperitoneal injection once a day for 5 days and perfused 3 days after the last BrdU injection.

The mice were deeply anesthetized and perfused with 1× phosphate-buffered saline (PBS) and ice-cold 4% paraformaldehyde (PFA) in PBS, pH 7.2. Brains were dissected and postfixed with 4% PFA overnight at 4°C. For cryoprotection, fixed brains were stored in 30% sucrose in PBS at 4°C. The brain was embedded in optimal cutting temperature compound (Tissue Tek, Sakura Finetek, 25608-930) and frozen at −80°C for cryosectioning. Frozen brains were serially sectioned with Leica CM 1850 or 1950 (Leica Microsystems, Wetzlar, Germany) in the coronal plane at 16 μm thickness. Every 15th sections (each slice 240 μm apart from the next) were serially mounted on individual Colorfrost Plus Microscope Slides (Fisher Scientific) in order from anterior to posterior and preserved at −20°C until use.

Animals for LacZ staining were perfused with PBS, and the dissected brains were postfixed with 2% PFA for 1.5 hr at 4°C. Cryosections were washed with PBS, and X-gal staining was developed at 37°C overnight in the staining solution (5 mM K₃Fe(CN)₆, 5 mM K₄Fe(CN)₆, 5 mM EGTA(ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid), 0.01% deoxycholate, 0.02% NP40, 2 mM MgC1₂, and 1 mg/ml X-gal). Sections were postfixed with 10% formalin at room temperature overnight, followed by counterstain with nuclear-fast red (H-3403, Vector Laboratories) at room temperature for 10 min before proceeding for dehydration (70%, 95%, 100% ethanol, xylene twice) and coverslipping with Mount-Quick (Ted Pella).

Cryosections were washed with PBS and incubated overnight at 4°C with primary antibodies diluted in blocking solution (10% Lamb serum and 0.3% Triton X-100). The following primary antibodies were used in this study: rabbit anti-Sox2 (1:1000; Abcam, ab92494, RRID:AB_10585428); rat anti-Sox2 (1:1000; Invitrogen, 14-9811-82, RRID:AB_11219471); rabbit anti-DCX (1:1000; Abcam, ab18723, RRID:AB_732011); chicken anti-GFAP (1:1000; Millipore, AB5541, RRID:AB_177521); rat anti-RFP (1:1000, Chromotek, 5f8-100, RRID:AB_2336064); chicken anti-S100B (1:500, Synapyic Systems, 287 006, RRID:AB_2713986); rat anti-BrdU (1:500, Abcam, ab6326, RRID:AB_305426) and mouse anti-BrdU (1:100, BD Biosciences, 347580, RRID:AB_400326); rabbit anti-GluR2/3 (1:100, Millipore, AB1506, RRID:AB_177521), rabbit anti-GluR2/3 (1:500, Epitomics, 1905), mouse anti-c-fos (1:500; Novus Biologicals, NBP2-50037, AB_2665387), rabbit anti-c-fos (1:1000, Synapyic Systems, 226 003, AB_2231974), rabbit anti-c-fos (1:100, Synapyic Systems, 226 008, RRID:AB_2891278). For staining of Sox2 and thymidine analogs, sections were heated in 10 mM citric acid pH 6.0 on boiling water bath for 15 min prior to blocking. After three washes in PBS, sections were incubated for 2 hr with corresponding secondary antibodies; goat anti-rat IgG (H+L) Alexa Fluor 546 (Invitrogen, A-11081, RRID:AB_2534125), donkey anti-rabbit IgG (H+L) Alexa Fluor 546 (Invitrogen, A-10040, RRID:AB_2534016), goat anti-chicken IgY (H+L) Alexa Fluor 546 (Invitrogen, A-11040, RRID:AB_2534097), goat anti-rabbit IgG (H+L) Alexa Fluor 633 (Invitrogen, A-21070; RRID: AB_2535731), donkey anti-mouse IgG (H+L) Alexa Fluor 647 (Invitrogen, A-31571, RRID:AB_162542), donkey anti-rabbit IgG (H+L) Alexa Fluor 488 (Invitrogen, A-21206,RRID:AB_2535792), and goat anti-rabbit IgG (H+L) Alexa Fluor 488 (Invitrogen, A-11008,RRID:AB_143165). Nucleus were stained by 4′,6-diamidino-2-phenylindole, dihydrochloride (Thermo Fisher, D1306). After a final rinse with PBS, sections were mounted on glass slides with Prolong gold antifade reagent (Thermo Fisher Scientific, P36930).

RNAscope in situ hybridization combined with immunofluorescence were performed using the RNAscope Multiplex Fluorescent V2 Assay (Advanced Cell Diagnostics) according to manufacturer’s protocol. Briefly, sections were dehydrated by ethanol and then treated with preheated co-detection target retrieval reagent (Advanced Cell Diagnostics). Subsequently, sections were incubated with primary antibody for rabbit anti-GluR2/3 (1:100, Millipore, AB1506, RRID:AB_177521) overnight at 4°C and post-fixed with 10% neutral buffered formalin. After three washes, the sections were treated with Protease Plus (Advanced Cell Diagnostics), and incubated with RNAscope probe for Shh, designed commercially by Advanced Cell Diagnostics (314361, RNAscope LS 2.5 Probe-Mm-Shh) for 2 hr at 40°C in HybEZ Oven (Advanced Cell Diagnostics). Detection of probe and antibody were performed using Tyramide Signal Amplification system with Opal 570 or 520 reagents (Akoya Biosciences, FP1488001KT and FP1487001KT). Expression of Shh in each mossy cell was evaluated by the number of Shh probe puncta in a GluR2/3-positive cell body. Shh mRNA signals in GluR2/3+ cells were counted in a series of sections spanning from anterior to posterior DG per mouse.

Images were acquired using a CSU-W1 spinning disk confocal microscope (Nikon) and Axioscan Z.1 (Carl Zeiss). NIH ImageJ (RRID:SCR_003070) was used for cell counting. Cell counting was performed in the DG of the hippocampus in a series of sections that were collected by serial mounting on 15 individual glass slides. Marker-positive cells in the indicated areas of the DG were counted in a series of sections on a single glass slide that contained DG at the same anatomical level between each group. The counts were then multiplied by the number of intervals between sections to estimate the total number of marker-positive cells in the DG. To quantify c-fos+ cell density in dentate hilus, c-fos+ cells were manually counted and normalized to the area size of hilus that are measured by ImageJ. For counting the total number of NSCs in the DG, radial NSCs were identified with a radial GFAP+ process extending from a Sox2+ nucleus in the subgranular zone of the DG. Persistence of NSC identity in each animal was calculated by dividing the total number of BrdU+ Sox2+ GFAP+ radial NSCs at 10 days or 4 weeks after BrdU administration by the total number of BrdU+ Sox2+ NSCs on the day of last BrdU administration (day 0) in the same group.

Statistical analyses were performed using either two-tailed unpaired t-test (for comparisons between two groups); one- or two-way analysis of variance with Tukey’s multiple comparison test (for multiple groups comparison) with Prism 9.0 software (GraphPad, RRID:SCR_002798). Differences were considered statistically significant at p < 0.05. Asterisks indicate significant differences (*<0.05; **<0.01, ***<0.001, ****<0.0001). No statistical methods were used to predetermine sample sizes. Sample sizes were based on previous studies using similar methods (Harris et al., 2021; Jessberger et al., 2007; Parent et al., 1997; Sierra et al., 2015; Yeh et al., 2018). Statistical analysis for specific experiments is shown in figure legends.