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. 2019 Apr 11;8:e42918. doi: 10.7554/eLife.42918

Suppressor of fused controls perinatal expansion and quiescence of future dentate adult neural stem cells

Hirofumi Noguchi 1, Jesse Garcia Castillo 1, Kinichi Nakashima 2, Samuel J Pleasure 3,✉
Editors: Francois Guillemot4, Jonathan A Cooper5
PMCID: PMC6459675  PMID: 30973324

Abstract

Adult hippocampal neurogenesis requires the quiescent neural stem cell (NSC) pool to persist lifelong. However, establishment and maintenance of quiescent NSC pools during development is not understood. Here, we show that Suppressor of Fused (Sufu) controls establishment of the quiescent NSC pool during mouse dentate gyrus (DG) development by regulating Sonic Hedgehog (Shh) signaling activity. Deletion of Sufu in NSCs early in DG development decreases Shh signaling activity leading to reduced proliferation of NSCs, resulting in a small quiescent NSC pool in adult mice. We found that putative adult NSCs proliferate and increase their numbers in the first postnatal week and subsequently enter a quiescent state towards the end of the first postnatal week. In the absence of Sufu, postnatal expansion of NSCs is compromised, and NSCs prematurely become quiescent. Thus, Sufu is required for Shh signaling activity ensuring expansion and proper transition of NSC pools to quiescent states during DG development.

Research organism: Mouse

Introduction

Newborn neurons are generated in two restricted regions of the adult rodent brain: the cortical subventricular zone (SVZ) and the dentate subgranular zone (SGZ) (Altman and Das, 1965; Eriksson et al., 1998; Kuhn et al., 1996; Lois and Alvarez-Buylla, 1993). Adult neurogenesis in the dentate gyrus (DG) has been implicated in hippocampal-dependent memory and learning (Deng et al., 2010). Newly generated neurons produced from neural stem cells (NSCs) residing in the SGZ are constantly added to the granule cell layer (GCL) and integrated into the existing hippocampal circuitry (Imayoshi et al., 2008). Persistence of adult hippocampal neurogenesis relies on the proper maintenance of NSCs even after development. However, little is known about the developmental programs governing the production and maintenance of long-lived NSCs.

Quiescence of NSCs during early development has been proposed to play a key mechanism for maintaining the NSC pool throughout life (Furutachi et al., 2013; Kawaguchi et al., 2013; Mira et al., 2010; Song et al., 2012). NSCs enter a quiescent state in a spatiotemporal manner during development, and this step is critical for ensuring the appropriate sized NSC pool for adult neurogenesis. At the beginning of forebrain development, NSCs are highly proliferative, but gradually lose proliferation competence with development and enter a quiescent state (Furutachi et al., 2015). The failure to transition to a quiescent state during development triggers continuous proliferation of NSCs and leads to premature exhaustion of the NSC pool (Furutachi et al., 2015). Furthermore, the NSC pool must be properly established since NSCs can only undergo a limited number of rounds of cell division prior to terminal differentiation. Live imaging of NSCs in adult DG and thymidine-analog based cell tracing analysis demonstrated that quiescent NSCs undergo a series of asymmetric divisions to produce neurons, and subsequently are consumed by symmetric differentiation into astrocytes or neurons (Calzolari et al., 2015; Encinas et al., 2011; Pilz et al., 2018). Indeed, adult neurogenesis and the NSC pool have been shown to decline with aging (Kuhn et al., 1996; Lugert et al., 2010), suggesting that for neurogenesis to persist throughout life, the size of putative quiescent NSC pool must be established during development before the NSCs transition to a quiescent state.

Long-lived NSCs in the DG originate from the ventral hippocampus at E17.5 (Li et al., 2013). These cells migrate along the longitudinal axis of the hippocampus from the temporal to septal poles and eventually settle in the ventral and dorsal DG. The initial production and maintenance of long-lived NSCs is dependent on Shh signaling and Shh ligands, produced by local neurons in the embryonic amygdala and the postnatal DG (Li et al., 2013). Blocking Shh signaling by deleting Smoothened (Smo) from responsive cells in the DG or ablation of Shh ligands from local neurons impairs the emergence of long-lived NSCs and results in diminishing the NSC pool (Han et al., 2008; Li et al., 2013). These findings highlight the significance of Shh signaling in production of the NSC pool during development. What is not clear yet from these studies is how Shh signaling activity is spatiotemporally regulated to ensure the expansion of the NSC pool during DG development and the role of Shh signaling in the transition of NSCs to a quiescent state.

Shh signaling is critical at early stages of embryonic brain development. Thus, complete ablation of Shh signaling activity by Smo deletion or the constitutive activation of Shh signaling by expressing an active Smo mutant (SmoM2) severely compromise the initial steps of DG development (Han et al., 2008). The embryonic nature of this phenotype prevents the further analysis of specific roles of Shh signaling in postnatal DG development, particularly in the production and maintenance of postnatal NSCs. To circumvent this, we are utilizing a Cre-loxP based system that allows spatiotemporal analysis of Shh signaling activity by genetic manipulation of the Shh signaling inhibitor, Suppressor of Fused (Sufu), a Gli-binding protein with an indispensable role in embryonic development. Conditional deletion of Sufu in a spatiotemporal manner allowed us to examine the role of Shh signaling in various aspects of NSC behavior during DG development. Our earlier studies showed that Sufu is important for the specification of NSC fate decision during cortical development via regulating Shh signaling activity (Yabut et al., 2015). In this report, we set out to determine the contribution of Sufu in regulating Shh signaling during DG development and how Sufu and Shh signaling are involved in the mechanisms governing the expansion of long-lived NSCs and their transition to the quiescent state during DG development. Intriguingly, we find that deletion of Sufu decreases Shh signaling in NSCs during DG development – this is in distinction to the neocortex where loss of Sufu increases Shh signaling. Long-lived NSCs expand in the early part of first postnatal week, but proliferation of these NSCs is impaired in the absence of Sufu, resulting in a decreased NSC pool in the adult DG. We also found that long-lived NSCs gradually become quiescent towards the end of the first postnatal week. However, Sufu deletion precociously triggers this transition to the quiescent state. Taken together, these results indicate that loss of Sufu during DG development decreases Shh signaling activity and impairs expansion of long-lived NSCs and the timely transition to a quiescent state during DG development.

Results

Deletion of Sufu in NSCs reduces Shh signaling during DG development

Shh ligands originate from amygdala neurons and the adjacent ventral dentate neuroepithelium to activate Shh signaling in ventral hippocampal NSCs (Li et al., 2013). These Shh-responding NSCs subsequently migrate to the dorsal DG and gradually accumulate between the hilus and GCL to form the SGZ postnatally (Li et al., 2013). Sufu is expressed in NSCs of the developing forebrain including presumptive DG cells (Yabut et al., 2015). We previously reported that deletion of Sufu in NSCs at early gestational stages (E10.5) severely disrupted the overall cytoarchitecture of the forebrain as a consequence of ectopic activation of Shh signaling (Yabut et al., 2015). To determine the effects of Sufu specifically in DG development, we used a hGFAP-Cre line to delete Sufu in NSCs at E13.5, before the initiation of DG development (E14.5) – we call these mice hGFAP-Sufu-KO. We first asked if deletion of Sufu increases Shh signaling activity. Expression of Gli1, a downstream target gene of Shh signaling, is strictly dependent on Shh signaling stimulation. Thus, we used Gli1-LacZ reporter mice (Bai et al., 2002), in which lacZ is expressed under the Gli1 promoter, to identify and characterize Shh-responding cells in the developing DG of hGFAP-Sufu-KO mice. Accordingly, we found abundant Gli1-lacZ + cells in the ventricular zone of the ventral hippocampus in Sufufl/fl;Gli1lacZ/+ mice (Figure 1A). Gli1-lacZ + cells were also present in the dorsal DG of Sufufl/fl;Gli1 lacZ/+mice at P0 and were enriched in the SGZ at P7 (Figure 1B–C and Figure 1—figure supplement 1). Surprisingly, we found a remarkable reduction of Gli1-LacZ + cells in ventral hippocampus of hGFAP-Sufu-KO;Gli1lacZ/+ mice at E15.5-P0, and small numbers detected throughout the anterior to posterior DG at P7. These data demonstrate that deletion of Sufu in NSCs decreases Shh signaling activity during DG development. This is distinct from the embryonic neocortex where deletion of Sufu increases Shh signaling activity (Yabut et al., 2015).

Figure 1. Deletion of Sufu decreases Hh-responding cells during DG development.

(A) Representative Gli1-LacZ staining images of sagittal brain sections in Sufufl/fl;Gli1lacZ/+and hGFAP-Sufu-KO;Gli1lacZ/+ mice at E15.5-P0. Magnified images of the black dashed-line boxes are shown to below of each image. (B) Representative Gli1-LacZ staining images of dorsal DG in sagittal sections of Sufufl/fl;Gli1lacZ/+ and hGFAP-Sufu-KO;Gli1lacZ/+ mice. (C) From anterior to posterior, four levels of coronal sections for Gli1-nLacZ staining at E17.5 are shown. Note that lacZ +cells are diminished in the DG of hGFAP-Sufu-KO;Gli1lacZ/+ mice from the beginning of DG development.

Figure 1.

Figure 1—figure supplement 1. Deletion of Sufu decreases Hh-responding cells during DG development.

Figure 1—figure supplement 1.

Representative Gli1-LacZ staining images of dorsal DG in sagittal sections of Sufufl/fl;Gli1lacZ/+ and hGFAP-Sufu-KO;Gli1lacZ/+ mice. Note that lacZ +cells are observed more in dorsal DG compared with ventral DG, and these cells are diminished in both DGs in hGFAP-Sufu-KO;Gli1lacZ/+ mice.
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Deletion of Sufu decreases proliferation of NSCs in developing DG

Shh signaling plays a pivotal role in establishing and maintaining the NSC pool to adulthood (Choe et al., 2015; Han et al., 2008; Li et al., 2013). Ablation of Shh signaling in NSCs by deleting Smo leads to a drastic reduction in NSC proliferation and results in the failure of SGZ establishment (Han et al., 2008; Li et al., 2013). Since hGFAP-Sufu-KO;Gli1lacZ/+ mice showed a reduction in Gli1 expression at the onset of DG development, we investigated if deletion of Sufu influences the proliferation capacity of NSCs particularly in the SGZ where Sox2 +cells form the NSC pool. During early postnatal DG development, NSCs migrate to the border between the hilus and GCL to form SGZ. The SGZ can be distinguished by accumulation of Sox2 +cells around P7 and is fully established by second postnatal week. Thus, we examined the number of Sox2 +cells in SGZ at P7 and P14. Although we found no difference in the number of Sox2 +cells between Sufufl/fl mice and hGFAP-Sufu-KO mice in the SGZ (Figure 2A and B), there was a significant reduction in Ki67 +proliferating cells in the P7 hGFAP-Sufu-KO mice (Figure 2C). In addition, there was a significant reduction in ratio of Ki67 +cells to Sox2 +NSCs in the SGZ of hGFAP-Sufu-KO mice (Figure 2D) indicating that very few Sox2 +cells retained their proliferative capacity. We next examined the number of Sox2 +cells and Ki67 proliferating cells in the SGZ of DGs at P14. In addition to the reduction in Ki67 +proliferating cells, the number of Sox2 +cells was significantly reduced in SGZ of hGFAP-Sufu-KO mice at P14 (Figure 2E–G). These data suggest that deletion of Sufu decreases NSC proliferation and number. We found that these phenotypes of Sufu-KO mice contrasts starkly with those observed in the hGFAP-Cre;SmoM2 mice, in which Shh signaling is constitutively active in NSCs. The DG of hGFAP-Cre;SmoM2 mice display abnormal morphology (Figure 2H) as previously reported (Han et al., 2008) and exhibit higher numbers of dying cells as marked by cleaved caspase 3 + cells (Figure 2—figure supplement 1A–C). Despite this, there was a significant increase in proliferating Ki67 +cells in the SGZ of hGFAP-Cre;SmoM2 compared to WT mice at P7 (Figure 2I and J). Accordingly, the proliferating population of Sox2 +cells was significantly higher than WT mice at P7 (Figure 2K). Taken together, these findings establish the crucial roles of Shh signaling in regulating NSC proliferation in the developing DG. Importantly, that deletion of Sufu resulted in downregulation, instead of ectopic activation, of Shh signaling in DG NSCs, indicates the distinct effects of Sufu in specific NSC subtypes and may involve a novel mechanism by which Sufu controls Shh signaling activity in DG NSCs.

Figure 2. Deletion of Sufu decreases proliferation of DG-NSCs.

(A) Representative immunofluorescence images of Sox2 (red) and Ki67 (green) in the DG of Sufufl/fl and hGFAP-Sufu-KO mice at P7. DNA is stained with DAPI (blue). Magnified images of the white dashed-line boxes are shown to the right of each image. (B,C) Quantification of Sox2+ (B) and Ki67+ (C) cells in the SGZ [Sufufl/fl, n = 5; hGFAP-Sufu-KO n = 7]. (D) The bar graph indicates the ratio of Ki67 +cells to Sox2 +NSCs in the SGZ [Sufufl/fl, n = 5; hGFAP-Sufu-KO n = 7]. (E) Representative immunofluorescence images of Sox2 (red) and Ki67 (green) in the DG of Sufufl/fl and hGFAP-Sufu-KO mice at P14. (F, G) Quantification of Sox2+ (F) and Ki67+ (G) cells in the SGZ [Sufufl/fl, n = 4; hGFAP-Sufu-KO n = 5]. (H) Representative immunofluorescence images of Sox2 (red), Ki67 (green) and DAPI (blue) in the DG of WT and hGFAP-Cre;SmoM2 mice at P7. Magnified images of the white dashed-line boxes are shown to the right of each image. (I,J) Quantification of Sox2+ (I) and Ki67+ (J) cells in the SGZ [WT, n = 4; hGFAP-Cre;SmoM2, n = 3]. (K) The bar graph indicates the ratio of Ki67 +cells to Sox2 +NSCs in the SGZ [WT, n = 4; hGFAP-Cre;SmoM2, n = 3]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01. Student’s t-test.

Figure 2—source data 1. Extended numerical data and statistical analysis for Figure 2—figure supplement 1.
elife-42918-fig2-data1.xlsx (41.4KB, xlsx)
DOI: 10.7554/eLife.42918.006

Figure 2.

Figure 2—figure supplement 1. Excessive Shh signaling activity leads to abnormal morphology of DGs and increased cell death.

Figure 2—figure supplement 1.

(A) Representative immunofluorescence images of Cleaved Caspase 3 (red) and DAPI (blue) in the DG of WT and hGFAP-Cre;SmoM2 mice at P7. (B, C) Quantification of Cleaved Caspase 3 + cells in the DGs (B) and SGZ (C) [n = 4 biological replicates per group]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01. Welch’s t-test.
Figure 2—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 2.
elife-42918-fig2-figsupp1-data1.xlsx (38.8KB, xlsx)
DOI: 10.7554/eLife.42918.007
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In the absence of Sufu, Gli1 function becomes responsible for proper proliferation of NSCs during DG development

Gli1, Gli2 and Gli3 are the main transcription factors that transduce Shh signaling to downstream targets. Gli3 mainly functions as a gene repressor (Hu et al., 2006; Litingtung et al., 2002; Persson et al., 2002; Wang et al., 2014), whereas Gli1 and Gli2 are responsible for activating target gene expression (Bai and Joyner, 2001; Park et al., 2000). Gli2 and Gli3 play major roles in regulating gene expression during the development, whereas Gli1 is largely dispensable (Bai et al., 2002; Park et al., 2000). Indeed, in the developing DG, we did not observe any differences in the number of Sox2 +cells and proliferating Sox2 +NSC in the SGZ between Sufufl/fl, Sufufl/fl;Gli1lacZ/+ and Sufufl/fl;Gli1lacZ/lacZ mice (Figure 3—figure supplement 1A–D). Therefore, we decided to use those genotypes as control. To our surprise, however, we found that deletion of one (hGFAP-Sufu-KO;Gli1lacZ/+) or both (hGFAP-Sufu-KO;Gli1lacZ/lacZ) Gli1 alleles in hGFAP-Sufu-KO mice led to a more profound phenotype. Overall, Ki67 +cells and the proliferating population of Sox2 +cells in the SGZ were significantly reduced in hGFAP-Sufu-KO;Gli1lacZ/+ and hGFAP-Sufu-KO;Gli1lacZ/lacZ mice compared with control at P7 (Figure 3A–C). In addition, we observed changes in NSC populations; hGFAP-Sufu-KO;Gli1lacZ/+ and hGFAP-Sufu-KO;Gli1lacZ/lacZ mice displayed a remarkable decline in Sox2 +cells in contrast with hGFAP-Sufu-KO mice at P7 (Figure 3D). These differences remained at P14 (Figure 3—figure supplement 2A–C). Given that the number of Sox2 +cells were comparable in hGFAP-Sufu-KO mice at P7 whereas significantly decreased in hGFAP-Sufu-KO;Gli1lacZ/+ mice, these findings indicate that deletion of Gli1 allele further impairs the proliferation of NSCs in GFAP-Sufu-KO mice.

Figure 3. Gli1 deletion increases the developmental defects in hGFAP-Cre Sufufl/fl mice.

(A) Representative immunofluorescence images of Sox2 (red), Ki67 (green) and DAPI (blue) in the DG of Control, hGFAP-Sufu-KO, hGFAP-Sufu-KO;Gli1lacZ/+, and hGFAP-Sufu-KO;Gli1lacZ/lacZ mice at P7. (B, C) Quantification of Ki67 +cells (B) and ratio of Ki67 +proliferating cell population in Sox2 +cells (C) in SGZ [Control, n = 8; hGFAP-Sufu-KO, n = 7; hGFAP-Sufu-KO;Gli1lacZ/+, n = 5; hGFAP-Sufu-KO;Gli1lacZ/lacZ, n = 5]. (D) Sox2 +cells counts in SGZ from each group [Control, n = 8; hGFAP-Sufu-KO, n = 7; hGFAP-Sufu-KO;Gli1lacZ/+, n = 5; hGFAP-Sufu-KO;Gli1lacZ/lacZ, n = 5]. (E) Representative immunofluorescence images of Tbr2 (red) and DAPI (blue) in the DG of Control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P7. (F) The number of Tbr2 +cells in DGs [Control, n = 7; hGFAP-Sufu-KO, n = 7; hGFAP-Sufu-KO;Gli1lacZ/+, n = 5]. (G) qRT-PCR analyses of Gli1, Gli2 and Gli3 expression in the P3 DGs of Sufufl/fl, Sufufl/fl;Gli1lacZ/+, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice [Sufufl/fl, n = 4; Sufufl/fl;Gli1lacZ/+, n = 4, hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01, ***p<0.001. ANOVA with Tukey post-hoc tests.

Figure 3—source data 1. Extended numerical data and statistical analysis for Figure 3.
elife-42918-fig3-data1.xlsx (43.4KB, xlsx)
DOI: 10.7554/eLife.42918.011

Figure 3.

Figure 3—figure supplement 1. Deletion of Gli1 does not affect cell proliferation.

Figure 3—figure supplement 1.

(A) Representative immunofluorescence images of Sox2 (red) and Ki67 (green) in the DG of Sufufl/fl, Sufufl/fl;Gli1lacZ/+, Sufufl/fl;Gli1lacZ/lacZ mice at P7. DNA is stained with DAPI (blue). (B,C) Quantification of Sox2+ (B) and Ki67+ (C) cells in the SGZ [Sufufl/fl, n = 4; Sufufl/fl;Gli1lacZ/+, n = 4; Sufufl/fl;Gli1lacZ/lacZ, n = 3]. (D) The bar graph indicates the ratio of Ki67 +cells to Sox2 +NSCs in DGs [Sufufl/fl, n = 4; Sufufl/fl;Gli1lacZ/+, n = 4; Sufufl/fl;Gli1lacZ/lacZ, n = 3]. Values represent mean ±SEM; ns: p>0.05. ANOVA with Tukey post-hoc tests.
Figure 3—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 3—figure supplement 1.
elife-42918-fig3-figsupp1-data1.xlsx (38.7KB, xlsx)
DOI: 10.7554/eLife.42918.012
Figure 3—figure supplement 2. Deletion of Sufu decreases proliferation of NSCs, but not neuronal precursors.

Figure 3—figure supplement 2.

(A) Representative immunofluorescence images of Sox2 (red) and Ki67 (green) in the DG of Control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P14. DNA is stained with DAPI (blue). (B,C) Quantification of Sox2+ (B) and Ki67+ (C) cells in the SGZ [Control, n = 6; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 5]. (D) Representative immunofluorescence images of Tbr2 (red) in the DG of Control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P14. (E) Tbr2 +cells counts in SGZ from each group [Control, n = 5; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. (F) Representative immunofluorescence images of Tbr2 (red) and Ki67 (green) in the DG of Control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P7. (G) The bar graph indicates the ratio of Ki67 +cells to Tbr2 +NSCs in DGs [Control, n = 4; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3]. (H) qRT-PCR analyses of Ptch1, N-Myc, Cyclin D1, and Cxcr4 expression in the DGs of Sufufl/fl, Sufufl/fl;Gli1lacZ/+, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P3 [Sufufl/fl, n = 4; Sufufl/fl;Gli1lacZ/+, n = 4, hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01, ***p<0.001. ANOVA with Tukey post-hoc tests.
Figure 3—figure supplement 2—source data 1. Extended numerical data and statistical analysis for Figure 3—figure supplement 2.
elife-42918-fig3-figsupp2-data1.xlsx (46.5KB, xlsx)
DOI: 10.7554/eLife.42918.013
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During the first postnatal week, NSCs give rise to neuronal precursor cells, which differentiate into granule neurons to form the GCL. Since neuronal precursor cells were produced from NSCs when they divide, we next examined if the reduction in proliferating Sox2 +cells correlated with a decrease in neuronal production. We found that Tbr2 +neuronal precursor cells, which was significantly reduced in hGFAP-Sufu-KO mice, was further reduced in hGFAP-Sufu-KO;Gli1lacZ/+ mice at P7 and P14 (Figure 3E,F and Figure 3—figure supplement 2D and E). Since neuronal precursors are also proliferative, we wondered if the reduced number of Tbr2 +cells in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice resulted from decreased proliferation of neuronal precursors. However, we found no difference in Ki67 +population of neuronal precursors between control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 3—figure supplement 2F and G). This suggests that deletion of Sufu does not impair the proliferation of neuronal precursors, and that impaired proliferation of NSCs leads to reduction of Tbr2 +cells in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. Together, these data suggest that deletion of Sufu impairs neuronal production in postnatal DG development, and this effect worsens when Gli1 expression is reduced.

The profound phenotypes of hGFAP-Sufu-KO;Gli1lacZ/+ mice compared with hGFAP-Sufu-KO suggest that deletion of Sufu increases the requirement for Gli1 function to regulate Shh signaling activity in DG development. To confirm this, we investigated the expression of Gli transcription factors transducing Shh signaling and their target genes. We extracted RNA from P3 DGs and examined the expression of Gli1, Gli2 and Gli3 by qPCR. As expected, Gli1 expression was reduced in the DG of hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 3G) and correlated with the reduction of Gli1-lacZ + cells (Figure 1). Interestingly, these mice also showed remarkable reduction in Gli2 and Gli3 expression (Figure 3G). The reduction of both Gli activators (Gli1 and Gli2) in hGFAP-Sufu-KO;Gli1lacZ/+ mice raise the possibility that reducing Gli1 in mice lacking Sufu further decreases Shh signaling activity in NSCs. This assumption is in line with previous reports of severe developmental defects when Gli2 levels are reduced in Gli1 homozygous null mutant mice (Bai et al., 2002; Park et al., 2000). To confirm reduced Shh signaling activity, we also checked the expression of Shh signaling target genes, Ptch1, N-myc, Cyclin D1 and Cxcr4 (Inaguma et al., 2015; Stecca and Ruiz i Altaba, 2005; Yin et al., 2019); Zheng et al., 2013), and found significant reductions in the expression of Shh signaling target genes in hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 3—figure supplement 2H). Taken together, these data suggest that Sufu deletion increases the dependency on Gli1 function during DG development, and that proliferation of NSCs is severely impaired when Gli1 expression is reduced in absence of Sufu.

Deletion of Sufu during DG development decreases NSC number and impairs adult neurogenesis

After development, NSCs in the DG are maintained until adulthood and produce neurons throughout life in the rodent brain. Given the widespread impairments in NSC numbers and proliferative capacity in the absence of Sufu, we next investigated the impact of these defects on adult neurogenesis. To label newborn neurons in adult mice, we administrated the thymidine analog 5-bromo-2’-deoxyuridine (BrdU) to 8 week old mice for 5 days and sacrificed the animal 3 days post-BrdU injection (Figure 4A). In hGFAP-Sufu-KO;Gli1lacZ/+ mice, the number of BrdU +newborn neurons, as identified by DCX, was significantly decreased (Figure 4B and C). Accordingly, there was a significant reduction of total DCX + cells and Tbr2 +cells in hGFAP-Sufu-KO;Gli1lacZ/+ mice compared with control mice (Figure 4D and Figure 4—figure supplement 1A and B). These data suggest that prenatal deletion of Sufu reduced adult neurogenesis. To determine if impaired neurogenesis is due to decreased differentiation of NSCs into neurons, we next compared the ratio of DCX + newborn neurons to BrdU labeled cells. However, there was no difference between control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 4E), suggesting that the observed defects in adult neurogenesis is not because of failure of NSCs to undergo neuronal differentiation.

Figure 4. Deletion of Sufu during DG development decreases qNSCs pool in adult DGs.

(A) Experimental scheme of BrdU injection. 8 week-old mice were injected with BrdU for 5 days and analyzed 3 days after last BrdU injection. (B) Representative immunofluorescence images for DCX (red) and DAPI (blue) in the DGs of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. Magnified images of DCX (red), BrdU (green) and DAPI (blue) are shown to the right of each image. (C, D) Quantification of BrdU+/DCX + cells (C) and DCX + cells (D) in SGZ [Control, n = 6; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 5]. (E) The bar graph indicates the ratio of DCX + cells to BrdU +cells in SGZ [n = 5 biological replicates per group]. (F) Representative immunofluorescence images for Sox2 (red), GFAP (cyan) and DAPI (blue) in the SGZ of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. (G) Quantification of Sox2+/GFAP +radial NSCs [Control, n = 6; hGFAP-Sufu-KO, n = 6; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. (H) The bar graph indicates the ratio of DCX + cells to Sox2+/GFAP +radial NSCs in SGZ [Control, n = 5; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, ***p<0.001. ANOVA with Tukey post-hoc tests.

Figure 4—source data 1. Extended numerical data and statistical analysis for Figure 4.
elife-42918-fig4-data1.xlsx (43.9KB, xlsx)
DOI: 10.7554/eLife.42918.016

Figure 4.

Figure 4—figure supplement 1. Deletion of Sufu during DG development decreases qNSCs pool in adult DGs.

Figure 4—figure supplement 1.

(A) Representative immunofluorescence images for Tbr2 (red) and DAPI (blue) in the DGs of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. (B) Quantification of Tbr2 +cells in SGZ [Control, n = 6; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. (C) Representative immunofluorescence images for Sox2 (red), Ki67 (green) and DAPI (blue) in the DGs of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. (D) The bar graph indicates the ratio of Ki67 +cells to Sox2 +cells in SGZ [Control, n = 6; hGFAP-Sufu-KO, n = 6; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. Values represent mean ±SEM; ns: p>0.05, *p<0.05. ANOVA with Tukey post-hoc tests.
Figure 4—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 4—figure supplement 1.
elife-42918-fig4-figsupp1-data1.xlsx (37.9KB, xlsx)
DOI: 10.7554/eLife.42918.017
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Impairments in adult neurogenesis may also arise from the failure of NSCs to undergo cell division. Therefore, we also examined the proliferation rate of adult NSCs by counting the proliferative Ki67 +population of Sox2 +cells. We found a comparable fraction of proliferating NSCs in the SGZ of DGs between control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 4—figure supplement 1C and D). This suggests that deletion of Sufu does not impair NSC proliferation at adult stages.

Adult NSCs are maintained in a quiescent state until stimulated to proliferate and produce neurons (Encinas et al., 2011; Mira et al., 2010). Since Sufu deletion reduced the number of Sox2 +cells in SGZ at P14, we investigated whether reduction of newborn neurons in hGFAP-Sufu-KO;Gli1lacZ/+ mice was due to a reduction in the production or maintenance of the quiescent NSC pool. These quiescent NSCs display a radial morphology with fiber extending to the molecular layer (Bignami and Dahl, 1974; Lugert et al., 2010; Rickmann et al., 1987; Sievers et al., 1992). To determine the number of quiescent NSCs in adult DGs, we counted the number of Sox2 +cells in SGZ, which have a GFAP +radial fiber. The number of Sox2+/GFAP +radial NSCs was significantly reduced in both hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice, with a greater reduction observed in hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 4F and G). Furthermore, we also calculated the ratio of DCX + cells to Sox2+/GFAP +radial NSCs to clarify the neurogenic competence of NSCs, and found no difference between control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 4H). This suggests that newborn neurons were produced from quiescent NSCs at comparable ratios and that the neurogenic competence was not impaired. Further, these findings supported our idea that decreased newborn neurons in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice results from reduction of NSC pool. These data suggest that deletion of Sufu in the developing DGs decreases the number of NSCs maintained through adulthood resulting in impaired adult neurogenesis.

Deletion of Sufu impairs proliferation and expansion of NSCs in the DG at early postnatal stages

Proliferation of NSCs in the first postnatal week is critical for producing and maintaining NSCs until adult stages (Youssef et al., 2018). Elimination of proliferating cells in the first postnatal week, but not at 2–3 weeks, severely impairs the size of the NSC pool in adult DG. We have previously shown that long-lived NSCs of the DG are composed of Hh-responding cells in the ventral hippocampus at E17.5 (Li et al., 2013). Using Gli1CreER/+::RosaAi14/+mice treated with Tamoxifen at E17.5, we labeled Hh-responding cells and traced their migration during the first postnatal week. We found that Ai14 +cells were sparsely localized in the dorsal DG and fimbriodentate junction (FDJ) at P0 (Figure 5A). However, the number of Ai14 +cells dramatically increased with development and accumulated in the border between hilus and GCL from P3 to P7. The number of Sox2 +cells in Ai14 +cells of ventrally derived NSCs was significantly increased from P0 to P3, and P3-P7, respectively (Figure 5B–D). Using Gli1CreER/+ mice, we also tested if inducing deletion of Sufu specifically in ventral NSCs decreases the number of NSCs. Gli1CreER/+;Sufufl/fl;Ai14 mice showed a significant reduction in Sox2+/Ai14 +cells and Sox2+/Ki67+/Ai14 +cells at P7 (Figure 5—figure supplement 1A–C), which is consistent with the results from GFAP-Cre mice (Figure 3). Together, these findings suggest that the first postnatal week is a critical period for long-lived NSC expansion.

Figure 5. Long-lived NSCs expand in first postnatal week, and loss of Sufu impairs its expansion.

(A) Fate tracing of Hh-responding cells at E17.5 in postnatal DG development. Gli1CreERT2/+ Ai14 mice were treated with tamoxifen at E17.5 and Ai14 +Hh responding cells are analyzed in postnatal first week (P0-7). Representative immunofluorescence images for Ai14 (red) and DAPI (blue) in dorsal and ventral hippocampus in sagittal sections. Note that Hh-responding cells labeled by Ai14 at E17.5 sparsely appear in DGs at P0 and subsequently accumulate in SGZ. (B) Representative immunofluorescence images for Ai14 (red), Sox2 (green) and DAPI (blue) in SGZ. (C,D) Quantification of Ai14+/Sox2 +cells in dorsal (C) and ventral (D) DGs [n = 3 biological replicates per group]. The number of long-lived NSCs, indicated as Ai14+/Sox2 +cells, are increased in postnatal first week. (E) BrdU was injected at P0, and pups were sacrificed 2 hr later. Representative immunofluorescence images for BrdU (green), Sox2 (red) and DAPI (blue) in dorsal and ventral DGs of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. (F) Schematic illustration of the dorsal and ventral DG at P0. d: dorsal, v: ventral, FDJ: fimbriodentate junction; SVZ: subventricular zone. The bar graph indicates the number of Sox2+/BrdU +cells in each region [Control, n = 6; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. (G) BrdU was injected at P3, and pups were sacrificed 2 hr later. Representative immunofluorescence images for BrdU (green), Sox2 (red) and DAPI (blue) in dorsal and ventral DGs of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice. (H) Quantification of Sox2+/BrdU +cells in dorsal and ventral DGs [Control, n = 5; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001. ANOVA with Tukey post-hoc tests.

Figure 5—source data 1. Extended numerical data and statistical analysis for Figure 5.
elife-42918-fig5-data1.xlsx (42.8KB, xlsx)
DOI: 10.7554/eLife.42918.020

Figure 5.

Figure 5—figure supplement 1. Prenatal deletion of Sufu in NSCs of ventral hippocampus leads to reduction of NSCs in the postnatal DGs.

Figure 5—figure supplement 1.

(A) Representative immunofluorescence images for Ai14 (red), Sox2 (green) and DAPI (blue) in the DGs of Gli1CreERT2/+;Sufufl/+;Ai14 and Gli1CreERT2/+;Sufufl/fl;Ai14 mice. (B,C) Quantification of Ai14+/Sox2 +cells (B) and Ai14+/Sox2+/Ki67 +cells (C) in DGs [n = 3 biological replicates per group]. Values represent mean ±SEM; *p<0.05. Student’s t-test.
Figure 5—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 5—figure supplement 1.
elife-42918-fig5-figsupp1-data1.xlsx (38.3KB, xlsx)
DOI: 10.7554/eLife.42918.021
Open in a new tab

In hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice, we found that the number of NSCs in adult DG was significantly reduced (Figure 4F and G). This might be attributed to impaired NSC expansion in the first postnatal week. To address this possibility, we labeled proliferating cells with BrdU at P0 or P3, and assessed the number of BrdU+/Sox2 +cells 2 hr post BrdU injection. At P0, there was no difference in the number of BrdU+/Sox2 +cells in both dorsal and ventral DGs between control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 5E,F). However, at P3, in dorsal, but not ventral DG, both hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice showed significant reduction in BrdU+/Sox2 +cells compared with control (Figure 5G and H). These findings showed that Sox2 +NSCs in the dorsal DG remain proliferative during the expansion period of long-lived NSCs. Taken together, these data suggest that Sufu deletion impairs the proliferation of NSCs at a critical expansion period resulting in reduced number of quiescent NSCs in the adult DG.

Deletion of Sufu leads to the premature transition of NSCs into quiescence during DG development

Our data show that Sufu deletion decreased NSC proliferation during the critical expansion period for long-lived NSCs, pointing to the likelihood that NSCs prematurely transitioned into a quiescent state. To test this, we utilized two thymidine analogs 5-Chloro-2-deoxyuridine (CldU) and 5-Iodo-2-deoxyuridine (IdU), and injected each thymidine analogs at different time points; CldU at P0, 3, 7 or 14 and IdU at 8 weeks old (Figure 6A and B). Because the thymidine analog is diluted as cells divide, cells that proliferated and stopped in the postnatal period will have detectable CldU, and therefore, when IdU is injected at adult stages, cells that became quiescent in developmental stages and then are reactivated in the adult will be double positive for CldU and IdU in adult stages. Double positive cells were observed more in mice injected with CldU at P3 and gradually decreased in groups injected with CldU at later postnatal stages (Figure 6C and D). Accordingly, the number of DCX+/CldU+/IdU +cells in newborn neurons was highest in groups injected at P3 (Figure 6E and F). Similarly, control mice injected with CldU at P3 had the highest number of CldU and IdU double positive cells at adult stages (Figure 6G and H). These data suggest that around P3-P7, NSCs in the control DG reduce their proliferation rate and become quiescent. However, in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice, the number of CldU and IdU double positive cells in CldU-injected groups at P3 or P7 was significantly decreased compared to control mice. Instead, detection of CldU and IdU double positive cells were significantly increased at P0 compared to control. The number of DCX + cells double labeled with CldU and IdU was also decreased in P3 or P7 CldU injected groups, whereas it was increased significantly in P0 CldU injected groups (Figure 6I). These data suggest that NSCs in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice prematurely exited the proliferative state from P0-3 instead of at P3-P7. To support this idea, we further clarified the timing of the transition towards quiescence by investigating the number of CldU+/Sox2+/GFAP +radial NSCs generated at each time point. Similar to our findings from the CldU and IdU double labeling experiments above, we found that the number of Sox2+/GFAP +radial NSCs labeled with CldU was decreased significantly in P3 and P7 CldU injected groups, whereas it was increased significantly in P0 CldU injected groups in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice (Figure 6—figure supplement 1A and B). In line with these observations, we also found that the Ki67 +proliferating population in Sox2 +NSCs was significantly reduced at P3, but not at P0 in hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice compared with control (Figure 6—figure supplement 2A–D). Comparable proportions of proliferating NSCs were found between three groups at P0 in the DG and FDJ where ventral-derived NSCs were localized at this time point (Figure 6—figure supplement 2A and B). However, we found that actively proliferating Sox2 +cells was significantly decreased in the SGZ of hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P3 and P7 (Figure 6—figure supplement 2C and D). Taken together, these data suggest that deletion of Sufu precociously reduces proliferation of NSCs and leads to premature transition to quiescent state and thus a smaller pool of quiescent NSCs in the adult dentate.

Figure 6. Deletion of Sufu prematurely induces the quiescent state transition.

(A) Schematic illustration for labeling NSCs that established quiescent state during DG development using two thymidine analogs. CldU is diluted when cell divides. Thus CldU amount goes down if the cell continuously proliferates during DG development. However, if the cell becomes quiescent state, CldU amount is maintained until adult then the cell can be double-labeled with CldU and IdU if the CldU-labeled quiescent cell starts proliferation and incorporate IdU at adult. (B) Experimental scheme of CldU and IdU injection. CldU was injected at P0, P3, P7 or P14, and then IdU was injected for 5 days at 8 weeks old. The mice were sacrificed at 3 days after last IdU injection. (C) Representative immunofluorescence images for CldU (green), IdU (red) and DAPI (blue) in the SGZ of animals injected CldU at different stages. White arrowheads indicate the CldU/IdU double-labeled cells. (D) Quantification of CldU+/IdU +cells in the SGZ [n = 3 biological replicates per group]. (E) Representative immunofluorescence images for CldU (green), IdU (red), DCX (cyan) and DAPI (blue) in the SGZ of animal injected CldU at P3. White arrowheads indicate the CldU+/IdU+/DCX + cells. (F) Quantification of CldU+/IdU+/DCX + cells in the SGZ [n = 3 biological replicates per group]. (G) Representative immunofluorescence images for CldU (green), IdU (red), DCX (cyan) and DAPI (blue) in the SGZ of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice injected CldU at P0. White arrowheads indicate the CldU+/IdU+/DCX + cells. (H,I) Quantification of CldU+/IdU +cells (H) and CldU+/IdU+/DCX + cells (I) in the SGZ [P0 CldU: Control, n = 7; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 6; P3 CldU: Control, n = 4; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3; P7 CldU: Control, n = 5; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01, ****p<0.0001. ANOVA with Tukey post-hoc tests.

Figure 6—source data 1 . Extended numerical data and statistical analysis for Figure 6.
elife-42918-fig6-data1.xlsx (43.3KB, xlsx)
DOI: 10.7554/eLife.42918.025

Figure 6.

Figure 6—figure supplement 1. Deletion of Sufu prematurely induces the quiescent state transition.

Figure 6—figure supplement 1.

(A) Representative immunofluorescence images for CldU (green), Sox2 (red), GFAP (cyan) and DAPI (blue) in the SGZ of animals injected CldU at different stages. White arrowheads indicate the CldU+/Sox2+/GFAP +radial NSCs. (F) Quantification of CldU+/Sox2+/GFAP +radial NSCs in the SGZ [P0 CldU: Control, n = 7; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 7; P3 CldU: Control, n = 4; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3; P7 CldU: Control, n = 5; hGFAP-Sufu-KO, n = 5; hGFAP-Sufu-KO;Gli1lacZ/+, n = 4]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, **p<0.01, ****p<0.0001. ANOVA with Tukey post-hoc tests.
Figure 6—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 6—figure supplement 1.
elife-42918-fig6-figsupp1-data1.xlsx (42.2KB, xlsx)
DOI: 10.7554/eLife.42918.026
Figure 6—figure supplement 2. Deletion of Sufu prematurely decreases the proliferation of NSCs during DG development.

Figure 6—figure supplement 2.

(A) Representative immunofluorescence images of Sox2 (red) and Ki67 (green) in the dorsal and ventral DG of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P0. DNA is stained with DAPI (blue). d: dorsal, v: ventral, FDJ: fimbriodentate junction; SVZ: subventricular zone. (B) The bar graph indicates the ratio of Ki67 +cells to Sox2 +NSCs in DGs [Control, n = 5; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3]. (C) Representative immunofluorescence images of Sox2 (red) and Ki67 (green) in SGZ of control, hGFAP-Sufu-KO and hGFAP-Sufu-KO;Gli1lacZ/+ mice at P3 and P7. (D) The bar graph indicates the ratio of Ki67 +cells to Sox2 +NSCs in DGs [P3: Control, n = 4; hGFAP-Sufu-KO, n = 4; hGFAP-Sufu-KO;Gli1lacZ/+, n = 3; P7: Control, n = 8; hGFAP-Sufu-KO, n = 7; hGFAP-Sufu-KO;Gli1lacZ/+, n = 5]. Values represent mean ±SEM; ns: p>0.05, *p<0.05, ***p<0.001. ANOVA with Tukey post-hoc tests.
Figure 6—figure supplement 2—source data 1. Extended numerical data and statistical analysis for Figure 6—figure supplement 2.
elife-42918-fig6-figsupp2-data1.xlsx (47.4KB, xlsx)
DOI: 10.7554/eLife.42918.027
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Discussion

Quiescence is key to maintaining the NSC pool and is critical for enabling lifelong neurogenesis, so understanding how actively dividing NSCs become quiescent during development is vitally important. Here, we demonstrated that long-lived NSCs dramatically expand in the first postnatal week before entering the quiescent state over several days. Sufu deletion impairs the ability of long-lived NSCs to expand in the first postnatal week, which results in the premature entry of NSCs into the quiescent state (Figure 7). This defect is a result of decreased Shh signaling activity as a result of Sufu deletion in NSCs.

Figure 7. Sufu is important for the perinatal expansion and quiescent state transition of dentate NSCs.

Figure 7.

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Schematic summary illustrating the role of Sufu and Shh signaling activity in initial production, expansion and quiescent state transition of NSCs during DG development. Long-lived NSCs are produced from Hh-responding cells in ventral hippocampus by Shh stimuli at E17.5 and migrate to dorsal and ventral DGs. This subset of NSCs expands in first postnatal week and subsequently become quiescent state. Sufu controls shh signaling activity in NSCs during DG development. Sufu deletion decreases the Shh signaling activity and leads to impaired expansion of long-lived NSC, resulting in premature quiescent state transition and small NSC pool in adult. On the other hand, complete ablation of Shh signaling activity by deleting Smo impairs initial production of long-lived NSCs at the beginning of DG development and compromises the establishment of neurogenic niche.

Surprisingly, our data showed that deletion of Sufu in GFAP-expressing NSCs decreased Shh signaling activity. Gli1-LacZ + cells were dramatically reduced in the embryonic ventral hippocampus and were abolished in the postnatal DG. Shh signaling is crucial for the initial production of dentate NSCs in the embryonic stage. Smo deletion in NSCs severely impairs initial NSC production and leads to decreased size of GCL (Han et al., 2008; Li et al., 2013). Although Sufu deletion impaired proliferation of NSCs in the first postnatal week, we still observed comparable number of proliferating NSCs at P0, indicating that initial NSC production at embryonic stages was not affected. Whether this process is dependent on Shh signaling is yet to be investigated, since we cannot exclude the possibility that very low levels of Shh signaling activity, likely undetectable by LacZ activity, are sufficient to produce the proper number of NSCs in embryonic stages.

Sufu is primarily known for its role as a negative regulator of Shh signaling pathway through several mechanisms - promoting the formation of Gli repressors, removing Gli1 from nucleus, and recruiting transcription repressors to Gli-target genes sites (Barnfield et al., 2005; Cheng and Bishop, 2002; Kise et al., 2009; Kogerman et al., 1999). However, recent studies provide evidence that Sufu may act as both a negative and positive regulator of Shh signaling under specific conditions. For example, in mouse embryonic fibroblast (MEF), Shh ligand binding triggers Sufu translocation to the nucleus with Gli1, where it binds to and facilitates cytoplasmic export of Gli3 repressor, thereby enhancing Shh signaling activity (Zhang et al., 2017). Furthermore, increasing the amount of Sufu added into Sufu-/- MEF compromises Hh-responsiveness in the absence of exogenous Shh, which is consistent with negative function of Sufu for Shh signaling. However, in the presence of exogenous Shh, Hh-responsiveness of Sufu-/- MEF is dramatically elevated with the addition of increasing levels of Sufu, an effect that was not observed when Ptch1 was added (Chen et al., 2009). These reports demonstrate that Sufu can act to enhance or maximize Shh signaling activity. Considering these roles, decreased Shh signaling activity in Sufu KO mice could result from the failure to maximize Shh signaling activity through mechanisms that likely involve stabilization of Gli transcription factors. Supporting this, our data showed that Gli1 and Gli2 expression were down-regulated in hGFAP-Sufu-KO;Gli1lacZ/+ mice in which expression of Shh signaling target genes were significantly reduced. Altogether, our data demonstrate multiple regulatory roles of Sufu in Shh signaling pathway that is dependent on cell type and context.

We found that deletion of Sufu increased the dependency of proper DG development on Gli1 function. Gli1 is not necessary for initial activation of Shh signaling (Bai et al., 2002; Park et al., 2000). Therefore, Gli1 deletion normally does not cause any significant developmental defects (Park et al., 2000). However, the phenotypes in hGFAP-Sufu-KO mice clearly worsen when combined with Gli1 deletion, suggesting that Gli1 function becomes necessary in the absence of Sufu. Previous studies show that Sufu is important for the stabilization of Gli2 and Gli3 proteins (Makino et al., 2015; Wang et al., 2010). Loss of Sufu results in diminishing Gli2 full-length activator. Sufu competitively binds to Gli2 and Gli3 with speckle-type POZ protein (Spop), which recruits ubiquitin ligases and degrades the full-length forms of Gli2 and Gli3 (Chen et al., 2009; Wang et al., 2010; Wen et al., 2010). These finding indicates that loss of Gli2 activators in the absence of Sufu increases the requirement for Gli1 function to maintain Shh signaling activity. Indeed, Gli1 is able to compensate for lost Gli2 activator function and rescue the developmental defects of Gli2 knockout mice (Bai and Joyner, 2001).

Expansion of long-lived NSCs must occur in the first postnatal week during which time ventral-derived NSCs dramatically increase their numbers in both the ventral and dorsal DG. Interestingly, Sufu deletion only impaired the proliferation of NSCs in the dorsal DG, but not ventral DG. This difference could be due to underlying molecular differences between NSCs and the surrounding cells residing in these regions. Lineage tracing of Shh expressing cell shows that neurons in the medial entorhinal cortex and hilar mossy cells function as the local source of Shh ligand for ventral derived NSCs, while hair mossy cells in the dorsal DG, but not ventral DG, are the source of Shh (Li et al., 2013). Conditionally removing Shh ligand from these local neurons abolishes signaling in Hh-responding cells and results in reduction of NSCs number, indicating that local Shh ligands are important for the activation of Shh signaling in NSCs after migration to the dorsal DGs (Li et al., 2013). Similar to mice lacking Shh, our data showed that deletion of Sufu abolished Hh-responding cells in postnatal DGs, followed by the reduction of proliferating NSCs in dorsal DGs. These findings indicate that dorsal and ventral NSCs use distinct developmental approaches to navigate the transition to produce long-lived NSCs. Dorsal NSCs must rely on Sufu to ensure optimal Shh signaling activity suitable for the expansion of long-lived NSCs during DG development, while NSCs in the ventral DG appear able to expand and become quiescent without Sufu.

Our data suggest that Shh signaling activity must be continuously maintained to promote NSC expansion and that eventual reduction in Shh signaling activity promotes NSCs to transition into the quiescent state. Supporting this, we found that Gli1-LacZ + cells were abundant in the ventral ventricular zone but progressively decreased after migration into the DG in the first postnatal week. Additionally, we found that constitutive activation of Shh signaling in NSCs, by conditional expression of SmoM2, increased proliferating NSCs and prevented transition into quiescence. These observations suggest that Sufu is important in sustaining NSC proliferation until NSCs begin to transition into a quiescent state. However, we also observed in SmoM2 mutants that subpopulations of NSCs become quiescent, indicating that reduction of Shh signaling activity alone is not sufficient to initiate quiescence and that other signaling mechanisms are involved in this process. Indeed, several extracellular factors can regulate the quiescent state of NSCs, such as Bone Morphogenetic Proteins (BMPs), Notch, and gamma-aminobutyric acid (GABA) (Kawaguchi et al., 2013; Mira et al., 2010; Song et al., 2012). These molecules are secreted by granule neurons, astrocytes, microglia, and interneurons in the DG and are likely sources of signals for migrating ventral-derived NSCs (Bonaguidi et al., 2011; Bond et al., 2014; Kawaguchi et al., 2013; Mira et al., 2010; Song et al., 2012; Yousef et al., 2015). The activity of these signaling pathways, and the simultaneous reduction in Shh signaling with development, may coordinately function to ensure successful transition of NSCs into the quiescent state during DG development.

Materials and methods

Key resources table.

Reagent type
(species) or
resource		 Designation Source or
reference		 Identifiers Additional
information
Genetic reagent
(M. musculus)		 Sufuflox/flox PMID: 20074523 RRID:
MGI:4840420 		Dr. Chi-Chung Hui
(University of Toronto)
Genetic reagent
(M. musculus)		 hGFAP-Cre PMID: 11668683 RRID:
MGI:2179048 		Jackson Laboratory
(Stock:004600)
Genetic reagent
(M. musculus)		 Gli1CreERT2/+ PMID: 15315762 RRID: MGI:3053957
Jackson Laboratory
(Stock:007913)
Genetic reagent
(M. musculus)		 Gli1LacZ/+ PMID: 12361967 RRID:
MGI:J:79392 		Jackson Laboratory
(Stock:008211)
Genetic reagent
(M. musculus)		 Rosa-Ai14 PMID: 20023653 RRID:
MGI:J:155793 		Jackson Laboratory
(Stock:007908)
Genetic reagent
(M. musculus)		 SmoM2-YFP PMID: 15107405 RRID:
MGI:3576373 		Jackson Laboratory
(Stock:005130)
Antibody Mouse monoclonal
anti-Ki67		 BD Biosciences RRID:
AB_396287 		IHC (1:500)
Antibody rabbit monoclonal
anti-Sox2		 Abcam RRID:
AB_10585428 		IHC (1:250)
Antibody Rabbit polyclonal
anti-Tbr2		 Abcom RRID:
AB_778267 		IHC(1:250)
Antibody rabbit polyclonal
anti-DCX		 Abcam RRID:
AB_732011 		IHC (1:1000)
Antibody rat monoclonal
anti-GFAP		 Zymed RRID:
AB_2532994 		IHC (1:500)
Antibody chickin polyclonal
anti-GFAP		 Millipore RRID:
AB_177521 		IHC (1:1000)
Antibody rat monoclonal
anti-RFP		 Chromotek RRID:
AB_2336064 		IHC (1:1000)
Antibody rat monoclonal
anti-BrdU		 Abcam RRID:
AB_305426 		IHC (1:500)
CldU detection
Antibody mouse
monoclonalanti-BrdU		 BD Biosciences RRID:
AB_400326 		IHC (1:100)
IdU detection
Antibody rabbit
monoclonalaniti-Cleaved
Caspase 3		 Cell Signaling RRID:
AB_2070042 		IHC (1:250)
Software ImageJ NIH RRID:
SCR_003070 		Cell counting
Software Prism 7 Graphpad RRID:
SCR_002798 		Statistuc analysis
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Animals

Mice carrying the floxed Sufu allele (Sufufl, ‘RRID:MGI:4840420’) were kindly provided by Dr. Chi-Chung Hui (University of Toronto) and were genotyped as described elsewhere (Pospisilik et al., 2010). The following mouse lines were obtained from Jackson Laboratory (Bar Harbor, Maine): Gli1CreERT2/+ (stock #007913, ‘RRID:MGI:3053957’), Gli1LacZ/+ (Stock #008211, ‘RRID:MGI:J:79392’), Rosa-AI14 (Stock #007908, ‘RRID:MGI:J:155793’), SmoM2 (Stock #005130, ‘RRID:MGI:3576373’), hGFAP-Cre (Stock #004600, ‘RRID:MGI:2179048’). Both male and female mice were analyzed with no distinction. All mice used in this study were maintained on a 12 hr light/dark cycle with free access to food and water. The day of vaginal plug was considered embryonic day 0.5. Mouse colonies were maintained at University of California San Francisco (UCSF) in accordance with National Institutes of Health and UCSF guidelines.

Tamoxifen and thymidine analog administration

Tamoxifen (Sigma) was dissolved in corn oil at 10 mg/ml. Pregnant mice were intraperitoneally administered 2 mg of tamoxifen with 27-gauge needles. For 5-Bromo-2’-deoxyuridine (BrdU) labeling, mice were subcutaneously (neonatal pups) or intraperitoneally (adult mice) injected with BrdU (Sigma) dissolved in saline (0.9% NaCl) at a dose of 50 mg/kg. For two thymidine analog labeling, 5-Chloro-2’-deoxyuridine (CldU, Sigma) dissolved in saline was subcutaneously injected into neonatal pups at a dose of 42.5 mg/kg followed by a single intraperitoneal injection of 57.5 mg/kg 5-Iodo-2’-deoxyuridine (IdU, Sigma) by for five days at 8 weeks old. IdU was dissolved in 0.015N NaOH and neutralized to pH 7.0 with 2N HCl before injection.

Tissue preparation

To prepare embryonic brain tissue, pregnant mice were sacrificed on the indicated developmental day, and embryos were perfused successively with phosphate-buffered saline (PBS) and ice-cold 4% paraformaldehyde (PFA) in PBS, pH 7.2. For preparation of postnatal and adult brains, pups and adult mice were deeply anesthetized before perfusion with 4% PFA in PBS. Brains were dissected and postfixed with 4% PFA in PBS 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 (OCT) compound (Tissue Tek, Sakura Finetek, 25608–930) and frozen at −80°C for cryosectioning. Frozen brains were serially sectioned with Leica CM 1850 (Leica Microsystems, Wetzlar, Germany) in the coronal or sagittal plane at 16 μm thickness. Every fifteenth sections were serially mounted on individual Colorfrost Plus Microscope Slides (Fisher Scientific) in order from anterior to posterior (coronal section) or medial to lateral (sagittal section), and preserved at −20°C until use.

LacZ staining and in situ hybridization

Animals for LacZ staining were perfused with 4% paraformaldehyde (PFA) and the dissected brains were postfixed with 4% PFA for 2 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, 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).

Immunohistochemistry

Cryosections were washed with PBS and blocked for 1 hr at room temperature with blocking solution (10% Lamb serum and 0.3% Triton X-100), and incubated overnight at 4°C with primary antibodies diluted in blocking solution. The following primary antibodies were used in this study: mouse anti-Ki67 (1:500; BD Biosciences, 550609, RRID:AB_396287); rabbit anti-Sox2 (1:1000; Abcam, ab92494, RRID:AB_10585428); rabbit anti-Tbr2 (1:1000; Abcam, ab23345, RRID:AB_778267); rabbit anti-DCX (1:1000; Abcam, ab18723, RRID:AB_732011); rat anti-GFAP (1:500; Zymed, 13–300, RRID:AB_2532994); chickin anti-GFAP (1:1000; Millipore, RRID:AB_177521); rat anti-RFP (1:1000, Chromotek, 5f8-100, RRID:AB_2336064); rat anti-BrdU (for BrdU or CldU detection) (1:500, Abcam, ab6326, RRID:AB_305426) and mouse anti-BrdU (for IdU detection)(1:100, BD Biosciences, 347580, RRID:AB_400326); rabbit anti-cleaved caspase 3 (1:250; Cell signaling #9661, RRID:AB_2070042). For staining of Ki67, Tbr2, Sox2 and thymidine analogs, sections were heated in 10 mM Citric acid pH 6.0 on boiling water bath for 10 min prior to blocking. For staining of Ai14 together with Ki67 and Sox2, sections were heated in 10 mM Citric acid pH 6.0 at 70 °C for 3 hr prior to blocking. After three washes in PBS, sections were incubated for 2 hr with corresponding secondary antibodies. After a final rinse with PBS, sections were mounted on glass slides with Prolong gold antifade reagent (Thermo Fisher Scientific).

Cell counting

Images were acquired using a Zeiss LSM 780 confocal microscope or Axio Scan Z.1 (Carl Zeiss). NIH ImageJ (RRID:SCR_003070) was used to count the cells. Brain sections were serially mounted on individual fifteen slides glasses. Cell counting was performed on every fifteenth sections containing DG at the same anatomical level between each group, and marker-positive cells were counted in the series of collected sections throughout the indicated areas in the DG. The total number of marker-positive cells in each indicated areas was obtained by multiplying the resultant counts by 15 (according to the interval between sections).

Real-time PCR

Total RNA was isolated using TRIzol Reagent (Thermo Fisher Scientific) and RNeasy Mini Kit (QIAGEN), according to the manufacturer’s instructions, and each sample was reverse-transcribed using a SuperScript IV cDNA Synthesis Kit (Invitrogen). Quantitative PCR reactions were performed using a SYBR Select Master Mix for CFX, and transcript expression was measured via CFX384 Touch Real-Time PCR Detection System (Bio-Rad). Expression levels of each gene were normalized to RNA polymerase II subunit A (polr2a) and calculated relative to the control. The primers used for this study are listed in Supplementary file 1.

Statistical analysis

At least three mice per group were analyzed. Statistical analyses were performed using either Student’s t-test or Welch’s t-test (for comparisons between two groups); one-way or two-way ANOVA with Tukey’s multiple comparison test (for multiple groups comparison) with Prism software (Graphpad, RRID:SCR_002798). All experiments were independently replicated at least three times. Differences were considered statistically significant at p<0.05. Asterisks indicate significant differences (*<0.05; **<0.01, ***<0.001, ****<0.0001).

Acknowledgements

We thank members of the SJP lab for helpful discussions, in particular OR Yabut and B Tran for technical help, suggestions, and helping to write this manuscript. This research was supported by NIH grant R01 NS075188 (SJP), Sasakawa Scientific Research Grant (HN), The Uehara Memorial Foundation (HN), JSPS Overseas Research Fellowships (HN).

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Samuel J Pleasure, Email: sam.pleasure@ucsf.edu.

Francois Guillemot, The Francis Crick Institute, United Kingdom.

Jonathan A Cooper, Fred Hutchinson Cancer Research Center, United States.

Funding Information

This paper was supported by the following grants:

  • Uehara Memorial Foundation to Hirofumi Noguchi.

  • Japan Society for the Promotion of Science Overseas Research Fellowship to Hirofumi Noguchi.

  • Japan Science Society Sasakawa Scientific Research Grant to Hirofumi Noguchi.

  • National Institutes of Health R01 NS075188 to Samuel J Pleasure.

Additional information

Competing interests

No competing interests declared.

Author contributions

Conceptualization, Resources, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing.

Conceptualization, Formal analysis, Validation, Investigation, Methodology, Writing—original draft, Writing—review and editing.

Formal analysis, Validation, Investigation.

Data curation, Supervision, Writing—review and editing.

Ethics

Animal experimentation: Mouse colonies were maintained at University of California San Francisco (UCSF) in accordance with National Institutes of Health and UCSF guidelines. Animal studies were approved by the Institutional Animal Care and Use Committee of UCSF (Protocol # AN176415-01A and AN165562-02A).

Additional files

Supplementary file 1. The primers for qPCR analysis.
elife-42918-supp1.xlsx (47.2KB, xlsx)
DOI: 10.7554/eLife.42918.029
Transparent reporting form
elife-42918-transrepform.pdf (337.5KB, pdf)
DOI: 10.7554/eLife.42918.030

Data availability

All data generated or analyzed in this study are included in the manuscript and supporting files.

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Decision letter

Editor: Francois Guillemot1

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Suppressor of Fused controls perinatal expansion and quiescence of future dentate adult neural stem cells" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom, Francois Guillemot, is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Jonathan Cooper as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

This is a timely manuscript that examines the mechanisms underpinning the expansion and entry into quiescence of dentate neural stem cells at early postnatal stages. The data supports a role of Sonic hedgehog (Shh) signaling regulated by Suppressor of Fused (Sufu) in this process. There is much interest in understanding how adult neural stem cells are generated during brain development and this manuscript has the potential to make a significant contribution in this field.

Summary:

The authors have previously shown that neural stem cells (NSCs) from the ventral hippocampus seed the subgranular zone of the postnatal dentate gyrus to form the adult NSC niche – a process that requires Shh signaling. This new work examines the effect of Sufu deletion on the establishment of the NSC pool in the mouse hippocampus.

Sufu is usually considered a repressor of the Shh pathway although it has also been shown to stimulate Shh signaling in certain contexts. This is the case here where the phenotypes arising from Sufu deletion resemble the phenotypes of a Smo floxed mutant mouse they analysed in their previous work (Li et al., 2013).

The authors conclude from a number of CldU/IdU incorporation and immunohistochemistry experiments that in wildtype animals, many NSCs enter into quiescence at the end of the first postnatal week. This is later than in the other adult NSC niche where NSCs lining the lateral ventricles enter quiescence at embryonic stages. They also find that decreasing Shh signaling by several mechanisms including Sufu deletion leads to precocious entry into quiescence, fewer NSCs in the hippocampus at 8 weeks of age and reduced neurogenesis at later time points.

The most novel aspect of this study is that it defines the precise timing of proliferation and entry into quiescence of NSCs during DG development and implicates downregulation of Shh signaling in promoting the transition of NSCs from proliferation to quiescence during development of the DG and thereby in determining the size of the adult NSC population.

Less novel is the establishment of a role of Sufu in enhancing Shh signaling, as Sufu is known to either stimulate or suppress Shh signaling depending on the cellular context. The implication of Shh signaling in the generation and proliferation of NSCs in the DG at early postnatal stages is also not novel as this was already shown in their previous paper (Li et al., 2013).

Overall, the data is well presented, the figures are of excellent quality and the paper well written, and the data mostly supports the conclusions reached (although not entirely, see below). However, there are several issues which will need to be resolved prior to publication.

Essential revisions:

1) The authors show that there is a large expansion of NSCs during normal development of the dentate gyrus between P0 to P7 (Figure 5A-D) and they claim that deletion of Sufu causes NSCs to precociously transition to a quiescent state (P0-3 instead of P3-7 normally), thus reducing the pool of quiescent NSCs in the adult (Figure 4F). However, Figure 2B clearly shows that there is no difference in the number of Sox2+ NSCs in the dentate gyrus of Sufu KO mice at P7. If the expansion of NSCs occurs from P0 to P7, then the decrease in NSC number should be apparent by P7. In agreement with this, the model presented in Figure 7 suggests that there are fewer NSCs (quiescent and dividing combined) in the Sufu KO at P3-7, but the data in Figure 2B clearly shows that there is no difference in the number of Sox2+ NSCs at P7. Along the same line, if there is an excessive proliferation of NSCs in SmoM2 mice (Figure 2G, H), why is their number unchanged at P7 (Figure 2F)? The authors should comment on these apparent contradictions and may need to amend their model.

2) The authors show in Figure 5G, H that Sufu KO causes reduced proliferation of NSCs in the dorsal but not ventral dentate gyrus at P3. However, in Figure 1C it appears as though most of the Shh-responsive cells are in the posterior/ventral dentate gyrus at P7, which is presumably true for P3 as well. Can the authors explain this discrepancy?

3) The authors show in Supplementary Figure 1 that Gli1 deletion has no effect on the number or proliferation of NSCs at P7 in the dentate gyrus and conclude that "Gli1 is largely dispensable [for forebrain development]". If this is true, then can the authors comment on why they think that a Gli1-lacZ reporter line is a good reporter of the level of Shh signaling required for development of the dentate gyrus (Figure 1)? In light of the fact that Sufu KO unexpectedly decreases Gli1-lacZ reporter expression, is there another way to confirm that Sufu KO decreases Shh signaling in the developing dentate gyrus? Related to this, the interpretation of the data on the regulation and function of Gli genes in wildtype and Sufu KO mice in Figure 3G is somewhat confusing and need to be clarified (subsection “In the absence of Sufu, Gli1 function becomes responsible for proper proliferation of NSCs during DG development”).

4) Important for the interpretation of the data is the assumption that DG development is not affected until the first postnatal week. The authors show that total Sox2+ cell numbers are unchanged. However, this appears to be a rather superficial read out and should be expanded by other measures (e.g., estimates of total neuronal and glial populations). If Sufu deletion affects the formation of the DG, all effects could be at least be partially explained by secondary effects. Using a conditional Cre line to test later stages of Sufu deletion (e.g., early postnatal) would be one way to address this concern. Related to this, the study focuses on the proliferation and quiescence of NSCs, yet much of the analysis is done by quantifying Sox2+ cells, which include both NSCs and the larger population of IPCs. Shh/Sufu might regulate the proliferation of IPCs as well as NSCs and it is unclear how much the changes in cell proliferation observed in loss and gain of function mutants concern specifically NSCs. The authors should use a more specific NSC marker or reassess their conclusions.

5) In Figure 4, the authors analysed neurogenesis in the adult DG and found reduced numbers of radial glia-like NSCs and reduced rate of neuron production, however it is unclear whether NSC proliferation is altered (percentage of NSCs that are Ki67+) and whether the total number of DCX+ cells produced decreases in adult Sufu KO mice.

6) The authors overinterpret their data regarding the role of Sufu in regulation of Shh signaling and NSC quiescence. The last sentence of the Introduction states that "Sufu (…) is important for ensuring (…) the timely transition to a quiescent state during DG development". However there is no evidence that Sufu is implicated in determining the timing of the transition to quiescence. The authors add at the beginning of the Discussion: "Thus Sufu modulates the timing of quiescence of NSCs in DG development by controlling NSCs expansion via modulation of Shh signaling activity". Again the paper does not provide evidence that Sufu normally modulates Shh activity and regulates the timing of quiescence of NSCs. Changes in Shh signaling and in timing of NSC quiescence are observed in Sufu mutant mice but there is no evidence that entry of NSCs in quiescence at the end of the first postnatal week in wildtype mice involves a modulation of Sufu function (i.e. another regulator might be normally involved in downregulating Shh signaling at that time). Please modify these interpretations of the data.

7) Why did the authors choose to do their analysis at P7 in Figure 2 and 3? The data from experiments in Figure 4-6 indicate that P7 is a transitional stage in which there is likely a mix of "developmental" and "adult-like" NSCs. Therefore, after going through Figure 4-6, it is unclear if the interpretation of the data at P7 is due to the shift in entry to quiescence or reduced proliferation of adult NSCs. As a result, the data in Figure 2 and 3 seems difficult to interpret and a bit out of place in the manuscript.

8) Overall, the dual thymidine analog experiment is difficult to interpret, because proliferation levels are different between genotypes at P0, 3, and 7, and the proliferation rates at 8 weeks are not presented (see comment #6). For example, if proliferation in adult DG decreased in Sufu KO animals, then IdU in the dual thymidine analog experiment would label fewer cells in Sufu KO animals, which would confound the interpretation of this experiment; the decrease in CldU+ IdU+ cells at P3 and P7 time points could be due to the altered transition of NSCs into quiescence or it could be due to the reduced proliferation in the adult or both. It might be simpler to interpret the results if you focus on CldU+ cells alone and look at how many CldU+ RGLs are generated at each time point in this experiment. On a similar note, most CldU+ IdU+ cells counted in Figure 6D and 6H are DCX+ (Figure 6F and 6I), which means that these data might reflect levels of proliferation and neurogenesis in the adult rather than altered entry into quiescence at postnatal stage. Additionally, the greater CldU+ IdU+ signals at P3 could indicate greater proliferation before P3 or greater quiescence after P3, or both. The gradual entry of NSCs into quiescence could be looked at in a simpler and more convincing fashion by examining the proportion of dividing (Ki67+) Sox2+ NSCs in wildtype mice from P0 to P7.

eLife. 2019 Apr 11;8:e42918. doi: 10.7554/eLife.42918.034

Author response

Essential revisions:

1) The authors show that there is a large expansion of NSCs during normal development of the dentate gyrus between P0 to P7 (Figure 5A-D) and they claim that deletion of Sufu causes NSCs to precociously transition to a quiescent state (P0-3 instead of P3-7 normally), thus reducing the pool of quiescent NSCs in the adult (Figure 4F). However, Figure 2B clearly shows that there is no difference in the number of Sox2+ NSCs in the dentate gyrus of Sufu KO mice at P7. If the expansion of NSCs occurs from P0 to P7, then the decrease in NSC number should be apparent by P7. In agreement with this, the model presented in Figure 7 suggests that there are fewer NSCs (quiescent and dividing combined) in the Sufu KO at P3-7, but the data in Figure 2B clearly shows that there is no difference in the number of Sox2+ NSCs at P7. Along the same line, if there is an excessive proliferation of NSCs in SmoM2 mice (Figure 2G, H), why is their number unchanged at P7 (Figure 2F)? The authors should comment on these apparent contradictions and may need to amend their model.

We understand the reviewers’ concerns. We have considered which stage is appropriate to examine the result of Sufu deletion. According to the data in Figure 6 (CldU and IdU), P7 is still a transitional stage in which NSCs shift to a quiescent state from a developmental proliferative state. Therefore, Sox2+ cells in the SGZ of P7 DGs would contain both states of NSCs, and there are still some expanding NSCs in the SGZ of P7 DGs. Given that the transition and expansion of NSCs are not completed yet at P7, it is difficult to determine the overall effects on the NSC pool in the Sufu-KO mice at this stage. Figure 6 demonstrated that the transition to quiescent state concludes by P14. Therefore, we have checked the number of Sox2+ cells at P14 as well (Figure 2 and Figure 3—figure supplement 2A-E). In P14 DGs, we found that the number of Sox2+ cells was significantly reduced in the SGZ of Sufu KO mice. We included this new data in Figures 2, 3 and Figure 3—figure supplement 2.

We also appreciate the helpful comments about Figure 7. We agree that Figure 7 did not comport well the data in the original Figure 2 and 3. So, we have now modified Figure 7, in which Sufu-KO mice have equal amount of total NSCs (quiescent and dividing combined) and less number of dividing NSCs at P7, whereas the number of total NSCs are less at P14 and Adult.

We also understand the reviewers’ concerns about SmoM2 mice. We further analyzed the lack of major increases in the absolute number of Sox2+ NSCs in SmoM2 DG at P7. As shown in Figure 2, there is an increase in proliferating cells as labeled by Ki67 at this stage, in contrast to Sufu-KO mice. Simultaneously, we also found increased cell death as indicated by the expression of cleaved caspase 3+ cells in the DGs of SmoM2 mice at P7 (Figure 2—figure supplement 1). Thus, despite the drastic increase in proliferating NSCs, many of these cells do not survive and this leads to the comparable numbers of Sox2+ NSCs in control and SmoM2 DG. We have now added this information in Figure 2—figure supplement 1 and the revised manuscript.

2) The authors show in Figure 5G, H that Sufu KO causes reduced proliferation of NSCs in the dorsal but not ventral dentate gyrus at P3. However, in Figure 1C it appears as though most of the Shh-responsive cells are in the posterior/ventral dentate gyrus at P7, which is presumably true for P3 as well. Can the authors explain this discrepancy?

We appreciate these comments because they point out the difficulty of visualizing the ventral DG in coronal sections. To help clarify specific DG regions, we have corrected the figures in this revised manuscript to include the areas we designated as posterior/ventral dentate gyrus (see Figure 1—figure supplement 1). Figure 1C of the original manuscript showed coronal sections, which unfortunately did not show images that included the ventral DG. We have now added images that include the entire DG and marked dorsal and ventral DG regions in Figure 1—figure supplement 1. In this figure we showed a higher number of Gli1-LacZ+ cells localized in the dorsal DG compared with the ventral DGs of Sufufl/fl;Gli1lacZ/+ mice. This supports our findings that there is a specific reduction in proliferating NSCs in the dorsal but not ventral DG.

3) The authors show in Supplementary Figure 1 that Gli1 deletion has no effect on the number or proliferation of NSCs at P7 in the dentate gyrus and conclude that "Gli1 is largely dispensable [for forebrain development]". If this is true, then can the authors comment on why they think that a Gli1-lacZ reporter line is a good reporter of the level of Shh signaling required for development of the dentate gyrus (Figure 1)? In light of the fact that Sufu KO unexpectedly decreases Gli1-lacZ reporter expression, is there another way to confirm that Sufu KO decreases Shh signaling in the developing dentate gyrus? Related to this, the interpretation of the data on the regulation and function of Gli genes in wildtype and Sufu KO mice in Figure 3G is somewhat confusing and need to be clarified (subsection “In the absence of Sufu, Gli1 function becomes responsible for proper 275 proliferation of NSCs during DG development”).

Thank you for helpful comments about function of Gli1 and Gli1-lacZ mice. We realized that we didn’t adequately discuss Gli1 function in activating Shh signaling and also why the Gli1 promoter has been utilized to reliably detect Shh signaling activity. Gli1 functions to support Gli2 transcription activator function, but is compensated for extensively by Gli2 (Bai et al., 2002; Park et al., 2000). In the absence of Gli1, other Gli activators such as Gli2 are able to compensate and activate Shh signaling target gene expression (Park et al., 2000). Therefore, deletion of Gli1 alone does not cause developmental defects. Instead, it produces severe developmental defects when combined with reduced Gli2 expression (Bai et al., 2002; Park et al., 2000). Nevertheless, Gli1 is a reliable and direct transcriptional target of Gli2. Thus, Gli1 expression is strictly induced upon Shh stimulation (Bai et al., 2002; Lee et al., 1997), making Gli1-LacZ mice a useful readout for Shh signaling activity (Ahn and Joyner, 2005; Balordi and Fishell, 2007; Ihrie et al., 2011; Machold et al., 2003). We also confirmed by in situ hybridization thatexpression of Ptch1, a downstream target of Shh signaling, is detected where LacZ+ cells are localized in Gli1-LacZ mice in the developing DG (see Author response image 1 below). We have added further explanation of the Gli1 reporter mice and the relationship of Gli1 and Gli2 in the revised manuscript.

Author response image 1.

Author response image 1.

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To further support our Gli1-LacZ findings and confirm that Shh signaling is reduced in Sufu-KO mice, we conducted qPCR to determine the expression levels of Shh signaling target genes. We tested the expression of Shh signaling target genes (Ptch1, N-Myc, CyclinD1 and Cxcr4) in P3 DGs. We found in Sufu-KO mice, significant reductions were observed in two Shh targets: CyclinD1 and Cxcr4 expression. On the other hand, in the Sufu-KO;Gli1lacZ/+ mice, significant reductions in the expression of Ptch1, N-Myc, CyclinD1 and Cxcr4 were observed (Figure 3—figure supplement 1H). Thus, the reduction in Gli1-LacZ activity in Sufu-KO;Gli1lacZ/+ simultaneously and accurately reflected a decrease in Shh signaling activity. We have now added new qPCR data in Figure 3G and Figure 3—figure supplement 2H of the revised manuscript.

In this study, although we could not address the exact mechanism, we believe that reduced expression of both Gli activators (Gli1 and Gli2) may partially explain the greater reductions of Shh signaling target genes expression and profound phenotypes in Sufu-KO;Gli1lacZ/+ mice. Indeed, it is demonstrated that reducing Gli2 level in Gli1 homozygous null mice (which develop normally), leads to multiple developmental defects and death soon after birth (Bai et al., 2002; Park et al., 2000). We have included this interpretation in the revised manuscript.

4) Important for the interpretation of the data is the assumption that DG development is not affected until the first postnatal week. The authors show that total Sox2+ cell numbers are unchanged. However, this appears to be a rather superficial read out and should be expanded by other measures (e.g., estimates of total neuronal and glial populations). If Sufu deletion affects the formation of the DG, all effects could be at least be partially explained by secondary effects. Using a conditional Cre line to test later stages of Sufu deletion (e.g., early postnatal) would be one way to address this concern. Related to this, the study focuses on the proliferation and quiescence of NSCs, yet much of the analysis is done by quantifying Sox2+ cells, which include both NSCs and the larger population of IPCs. Shh/Sufu might regulate the proliferation of IPCs as well as NSCs and it is unclear how much the changes in cell proliferation observed in loss and gain of function mutants concern specifically NSCs. The authors should use a more specific NSC marker or reassess their conclusions.

We understand the concern that Sufu deletion by hGFAP-Cre may impair DG development before postnatal stages. In the revised manuscript, we show that there is no difference in the proliferation of NSCs at P0 (Figure 5E and Figure 6—figure supplement 2A and B). Therefore, we believe that prenatal DG development is not impaired by deletion of Sufu. We also have tested the effect of Sufu deletion temporally using Gli1CreER/+ mice by specifically inducing Sufu deletion at E17.5 (Gli1CreER/+;Sufu-KO; Ai14). In this mouse, Cre recombination (indicated by expression of Ai14) is induced in NSCs in the ventral hippocampus, which later migrate to the DG and become adult NSCs as we showed in Figure 5. Thus, we believe that Sufu deletion is induced specifically in NSCs in this mouse model. This would minimize the secondary effects and potential developmental abnormalities at prenatal stages. We analyzed these mice at P7 and found that the number of Sox2+/Ai14+ cells and Sox2+/Ki67+/Ai14+cells were significantly reduced in Gli1CreER/+;Sufufl/fl;Ai14 mice compared with Gli1CreER/+;Sufufl/+;Ai14. These findings are consistent with the phenotypes observed in the hGFAP-Cre mice. We have now added these data to Figure 5—figure supplement 1A-C.

As suggested by the reviewers, we investigated the effects of Sufu deletion on IPC production and proliferation in Sufu-KO mice. We found that the number of Tbr2+ IPCs was significantly reduced in Sufu-KO mice (Figure 3E, F and Figure 3—figure supplement 2D-G). Further assessments showed that the reduction in Tbr2+ cells in Sufu-KO mice is not due to impaired or reduced proliferation of IPCs since we found no differences in the Ki67+ proliferating cell population in Tbr2+ IPCs between control and Sufu-KO mice Figure 3—figure supplement 2F. These data suggest that deletion of Sufu does not affect the proliferation of IPC. Rather, our analyses show that deficits in IPC numbers were due to the failure of NSCs to generate this cell population. Thus, we believe that reduction of the proliferating cell population in Sox2+ in Sufu-KO mice reflects reduction of NSC proliferation. We have added these results in Figure 3 and Figure 3—figure supplement 2.

5) In Figure 4, the authors analysed neurogenesis in the adult DG and found reduced numbers of radial glia-like NSCs and reduced rate of neuron production, however it is unclear whether NSC proliferation is altered (percentage of NSCs that are Ki67+) and whether the total number of DCX+ cells produced decreases in adult Sufu KO mice.

Thank you for helpful suggestions. We have now added data examining the percentage of Ki67+ cells in Sox2+ NSCs in Figure 4—figure supplement 1C and D, in which there is no difference in the proliferating cell population of NSCs in adult DG between control, Sufu-KO and Sufu-KO;Gli1lacZ/+ mice. We have also investigated the number of total DCX+ cells and Tbr2+ cells (Figure 4D and Figure 4—figure supplement 1A and B). We found that DCX+ cells and Tbr2+ cells were significantly reduced in Sufu-KO;Gli1lacZ/+ mice. We have added these data in Figure 4 and Figure 4—figure supplement 1.

6) The authors overinterpret their data regarding the role of Sufu in regulation of Shh signaling and NSC quiescence. The last sentence of the Introduction states that "Sufu (…) is important for ensuring (…) the timely transition to a quiescent state during DG development". However there is no evidence that Sufu is implicated in determining the timing of the transition to quiescence. The authors add at the beginning of the Discussion: "Thus Sufu modulates the timing of quiescence of NSCs in DG development by controlling NSCs expansion via modulation of Shh signaling activity". Again the paper does not provide evidence that Sufu normally modulates Shh activity and regulates the timing of quiescence of NSCs. Changes in Shh signaling and in timing of NSC quiescence are observed in Sufu mutant mice but there is no evidence that entry of NSCs in quiescence at the end of the first postnatal week in wildtype mice involves a modulation of Sufu function (i.e. another regulator might be normally involved in downregulating Shh signaling at that time). Please modify these interpretations of the data.

We have modified our statements to clearly reflect our interpretations of how Sufu plays a role in regulating NSC quiescence. As the reviewers point out, there is no direct evidence that Sufu regulates the timing. Additionally, we have included in the Discussion other factors previously reported to play a role in regulating the production and timing of qNSCs in the DG. Whether or not Sufu functions in concert with these factors is not yet known and could be an interesting future direction.

7) Why did the authors choose to do their analysis at P7 in Figure 2 and 3? The data from experiments in Figure 4-6 indicate that P7 is a transitional stage in which there is likely a mix of "developmental" and "adult-like" NSCs. Therefore, after going through Figure 4-6, it is unclear if the interpretation of the data at P7 is due to the shift in entry to quiescence or reduced proliferation of adult NSCs. As a result, the data in Figure 2 and 3 seems difficult to interpret and a bit out of place in the manuscript.

This is a reasonable point. P7 is not necessarily the optimal timing to test the effects of Sufu deletion since it is such a transitional time in dentate formation; to get a more full picture it would be helpful to examine a slightly more advanced stage. Therefore, we have also analyzed the DG at P14, in which transition of "developmental" and "adult-like" NSCs seems largely complete. At this time point, we found that Sox2+ cells were significantly reduced in the DGs of Sufu-KO mice. Thus, P14 is an appropriate time to detect any defects arising from impaired expansion of the NSC pool. We have added these P14 data in the revised manuscript (Figure 2 and Figure 3—figure supplement 2).

8) Overall, the dual thymidine analog experiment is difficult to interpret, because proliferation levels are different between genotypes at P0, 3, and 7, and the proliferation rates at 8 weeks are not presented (see comment #6). For example, if proliferation in adult DG decreased in Sufu KO animals, then IdU in the dual thymidine analog experiment would label fewer cells in Sufu KO animals, which would confound the interpretation of this experiment; the decrease in CldU+ IdU+ cells at P3 and P7 time points could be due to the altered transition of NSCs into quiescence or it could be due to the reduced proliferation in the adult or both. It might be simpler to interpret the results if you focus on CldU+ cells alone and look at how many CldU+ RGLs are generated at each time point in this experiment. On a similar note, most CldU+ IdU+ cells counted in Figure 6D and 6H are DCX+ (Figure 6F and 6I), which means that these data might reflect levels of proliferation and neurogenesis in the adult rather than altered entry into quiescence at postnatal stage. Additionally, the greater CldU+ IdU+ signals at P3 could indicate greater proliferation before P3 or greater quiescence after P3, or both. The gradual entry of NSCs into quiescence could be looked at in a simpler and more convincing fashion by examining the proportion of dividing (Ki67+) Sox2+ NSCs in wildtype mice from P0 to P7.

We agree that decreased neurogenesis might make the analysis using CldU and IdU more confusing. We have also analyzed how many CldU+ RGLs are generated at each time point in this experiment (Figure 6—figure supplement 1). We counted the number of CldU+/Sox2+/GFAP+ radial NSCs and found a similar trend to what we observed in CldU/IdU experiments; the number of CldU+/Sox2+/GFAP+ radial NSCs was decreased significantly in P3 and P7 CldU-injected groups, whereas it was increased significantly in P0 CldU injected groups in Sufu-KO and Sufu-KO;Gli1lacZ/+ mice.

Following the reviewers’ suggestion, we also examined the proportion of dividing (Ki67+) Sox2+ NSCs from P0 to P7. We found that the proliferating cell population was comparable in P0 DG. However, it was decreased at P3 and P7 in the SGZ of Sufu-KO and Sufu-KO;Gli1lacZ/+ mice. These data indicate that deletion of Sufu prematurely decreases the proliferation of NSCs. We have added those results in Figure 6—figure supplement 2.

Associated Data

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    Supplementary Materials

    Figure 2—source data 1. Extended numerical data and statistical analysis for Figure 2—figure supplement 1.
    elife-42918-fig2-data1.xlsx (41.4KB, xlsx)
    DOI: 10.7554/eLife.42918.006
    Figure 2—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 2.
    elife-42918-fig2-figsupp1-data1.xlsx (38.8KB, xlsx)
    DOI: 10.7554/eLife.42918.007
    Figure 3—source data 1. Extended numerical data and statistical analysis for Figure 3.
    elife-42918-fig3-data1.xlsx (43.4KB, xlsx)
    DOI: 10.7554/eLife.42918.011
    Figure 3—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 3—figure supplement 1.
    elife-42918-fig3-figsupp1-data1.xlsx (38.7KB, xlsx)
    DOI: 10.7554/eLife.42918.012
    Figure 3—figure supplement 2—source data 1. Extended numerical data and statistical analysis for Figure 3—figure supplement 2.
    elife-42918-fig3-figsupp2-data1.xlsx (46.5KB, xlsx)
    DOI: 10.7554/eLife.42918.013
    Figure 4—source data 1. Extended numerical data and statistical analysis for Figure 4.
    elife-42918-fig4-data1.xlsx (43.9KB, xlsx)
    DOI: 10.7554/eLife.42918.016
    Figure 4—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 4—figure supplement 1.
    elife-42918-fig4-figsupp1-data1.xlsx (37.9KB, xlsx)
    DOI: 10.7554/eLife.42918.017
    Figure 5—source data 1. Extended numerical data and statistical analysis for Figure 5.
    elife-42918-fig5-data1.xlsx (42.8KB, xlsx)
    DOI: 10.7554/eLife.42918.020
    Figure 5—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 5—figure supplement 1.
    elife-42918-fig5-figsupp1-data1.xlsx (38.3KB, xlsx)
    DOI: 10.7554/eLife.42918.021
    Figure 6—source data 1 . Extended numerical data and statistical analysis for Figure 6.
    elife-42918-fig6-data1.xlsx (43.3KB, xlsx)
    DOI: 10.7554/eLife.42918.025
    Figure 6—figure supplement 1—source data 1. Extended numerical data and statistical analysis for Figure 6—figure supplement 1.
    elife-42918-fig6-figsupp1-data1.xlsx (42.2KB, xlsx)
    DOI: 10.7554/eLife.42918.026
    Figure 6—figure supplement 2—source data 1. Extended numerical data and statistical analysis for Figure 6—figure supplement 2.
    elife-42918-fig6-figsupp2-data1.xlsx (47.4KB, xlsx)
    DOI: 10.7554/eLife.42918.027
    Supplementary file 1. The primers for qPCR analysis.
    elife-42918-supp1.xlsx (47.2KB, xlsx)
    DOI: 10.7554/eLife.42918.029
    Transparent reporting form
    elife-42918-transrepform.pdf (337.5KB, pdf)
    DOI: 10.7554/eLife.42918.030

    Data Availability Statement

    All data generated or analyzed in this study are included in the manuscript and supporting files.

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