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. 2020 Feb 3;39(5):e101679. doi: 10.15252/embj.2019101679

Ependyma‐expressed CCN1 restricts the size of the neural stem cell pool in the adult ventricular‐subventricular zone

Jun Wu 1,2,3,4,†, Wen‐Jia Tian 2,3,4,5,†, Yang Liu 6,7,8, Huanhuan J Wang 2,3,4,5, Jiangli Zheng 1,2,3,4, Xin Wang 7,9, Han Pan 9, Ji Li 1, Junyu Luo 6, Xuerui Yang 7,9, Lester F Lau 10, H Troy Ghashghaei 11, Qin Shen 3,4,12,✉
PMCID: PMC7049812  PMID: 32009252

Abstract

Adult neural stem cells (NSCs) reside in specialized niches, which hold a balanced number of NSCs, their progeny, and other cells. How niche capacity is regulated to contain a specific number of NSCs remains unclear. Here, we show that ependyma‐derived matricellular protein CCN1 (cellular communication network factor 1) negatively regulates niche capacity and NSC number in the adult ventricular–subventricular zone (V‐SVZ). Adult ependyma‐specific deletion of Ccn1 transiently enhanced NSC proliferation and reduced neuronal differentiation in mice, increasing the numbers of NSCs and NSC units. Although proliferation of NSCs and neurogenesis seen in Ccn1 knockout mice eventually returned to normal, the expanded NSC pool was maintained in the V‐SVZ until old age. Inhibition of EGFR signaling prevented expansion of the NSC population observed in CCN1 deficient mice. Thus, ependyma‐derived CCN1 restricts NSC expansion in the adult brain to maintain the proper niche capacity of the V‐SVZ.

Keywords: CCN1, EGFR, ependymal cells, neural stem cells, niche

Subject Categories: Neuroscience, Regenerative Medicine

Secretion of the matricellular protein CCN1 from ependymal cells regulates adult neural stem cell (NSCs) expansion but also the capacity of stem cell niches for holding a specific number of NSCs and their progeny.

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Introduction

In the adult rodent brain, neurogenesis occurs mainly in two regions: the ventricular–subventricular zone (V‐SVZ) lining the lateral walls of the lateral ventricles and the subgranular zone (SGZ) in the hippocampal dentate gyrus (Kriegstein & Alvarez‐Buylla, 2009; Suh et al, 2009; Bond et al, 2015). In both regions, neural stem cells correspond to a subpopulation of astroglial cells and continue to generate neurons and glia throughout life (Kriegstein & Alvarez‐Buylla, 2009). In the adult V‐SVZ, B1 NSCs are surrounded and anchored by an array of ependymal cells, forming a pinwheel structure along the planar plane of the ventricular surface (Mirzadeh et al, 2008; Shen et al, 2008; Shook et al, 2012). One or more B1 NSCs are contained in the center of the pinwheel, separated by the surrounding ependymal cells (Mirzadeh et al, 2008), establishing NSC units that are spatially dispersed in the V‐SVZ. B1 cells are largely quiescent NSCs (qNSC) (Codega et al, 2014), characterized by lack of EGFR and the presence of the membrane protein vascular cell adhesion molecule 1 (VCAM1) on the apical endfeet, which directly contact the ependymal cells and protrude into the CSF (Kokovay et al, 2012; Llorens‐Bobadilla et al, 2015; Basak et al, 2018). Upon activation, a small percent of active NSCs (aNSC) self‐renew to maintain the stem cell pool, some of which return to quiescence, while the majority progress through a lineage from transient‐amplifying cells (C cells) to neuroblasts (A cells), generating olfactory bulb (OB) neurons after several rounds of division (Doetsch et al, 1999a; Calzolari et al, 2015; Basak et al, 2018; Obernier et al, 2018).

Despite long‐term self‐renewal, the abundance of NSCs decreases progressively during aging, which is attributed to consuming differentiation divisions and age‐related changes in cell‐intrinsic and niche‐related factors (Conover & Shook, 2011a; Shook et al, 2012; Capilla‐Gonzalez et al, 2014; Obernier et al, 2018). To compensate for the loss of NSCs, strategies to stimulate self‐renewal and proliferation of NSC are often exploited. However, expanding NSC pool by over‐activation of NSCs would compromise the long‐term potential of neurogenesis and lead to premature exhaustion of stem cells if the activated stem cells are not allowed to return to quiescence (Kippin et al, 2005; Mira et al, 2010; Furutachi et al, 2013; Jones et al, 2015; Urban et al, 2016; Neuberger et al, 2017; Zhou et al, 2018). In addition, in a closed stem cell niche, the amount of stem cells is limited by the physical constrain of niche architecture. For example, in the embryonic epidermis, active proliferation of progenitor cells leads to crowding, which subsequently triggers delamination and differentiation to maintain a stabilized niche size (Chacon‐Martinez et al, 2018). In the small intestine, slow‐dividing and label‐retaining stem cells are held in the specific locations at a constant amount under homeostatic condition (Barker et al, 2007; Takeda et al, 2011). Hence in a given stem cell niche, the number of quiescent stem cells and the self‐renewal potential of activated stem cells determine the ability of the niche to sustain a life‐long supply of newborn cells. Accumulating evidence from lineage tracing and single‐cell sequencing studies indicates that in the NSC niches, there is bidirectional transition between the states of qNSCs and aNSCs (Urban et al, 2016; Basak et al, 2018; Kester & van Oudenaarden, 2018; Obernier et al, 2018). However, the cellular and molecular mechanisms controlling the capacity of the NSC niche to hold a balanced number of qNSCs and aNSCs remain unclear.

Ependymal cells, occupying the majority of the ventricular surface area, are the closest neighbors of B1 cells in the V‐SVZ NSC niche. Although it has been shown that ependymal cell‐derived secreted factors, such as PEDF and Noggin, regulate NSC self‐renewal and neurogenic potential (Lim et al, 2000; Peretto et al, 2004; Ramirez‐Castillejo et al, 2006), how ependymal cells control the NSC niche and stem cell behavior remains largely unknown.

Cellular communication network factor 1 (Ccn1), originally named as Cyr61, was first identified as an immediate‐early gene. It belongs to the CCN family of genes encoding secreted proteins with a high degree of sequence homology and conserved tetramodular organization (Obrien et al, 1990; Perbal et al, 2018). CCN1 has been shown to promote growth factor‐induced proliferative responses through interacting with various cell surface integrins (Obrien et al, 1990; Babic et al, 1998). Upon secretion, CCN1 binds to the cell membrane and extracellular matrix (ECM) to elicit diverse cellular functions, including cell adhesion, migration, proliferation, survival, apoptosis, differentiation senescence, and tumor invasion (Chen & Lau, 2009). Although it has been shown that Ccn1 is expressed in the developing and adult brain, and high level of CCN1 correlates with a poorer prognosis in glioblastoma patients (Haseley et al, 2012; Ishida et al, 2015; Otani et al, 2017), the expression pattern and function of CCN1 in the adult NSC niche have not been studied yet.

In this study, we investigated the cellular and molecular relationship between NSCs and ependymal cells. We identified CCN1 as an ependyma‐specific niche factor, and our results imply that ependymal cells restrict niche capacity and NSC pool expansion in the adult V‐SVZ.

Results

CCN1 is specifically expressed in ependymal cells in the adult V‐SVZ

We characterized the expression pattern of CCN1 in the adult V‐SVZ by immunostaining on forebrain coronal sections and V‐SVZ whole‐mounts from 8‐ to 12‐week mice. Two different antibodies revealed similar patterns. In coronal sections, CCN1 was detected selectively in the apical cell layer lining the lateral ventricle (Fig 1A). From the en face view of the V‐SVZ whole‐mounts prepared from the lateral walls of the lateral ventricles, CCN1 was found in S100β+ cells with a large apical surface, indicating CCN1 is expressed in ependymal cells (Figs 1B and EV1A). To determine whether CCN1 was also expressed in apical B1 cells, which are embedded in the ependymal layer, we labeled B1 cells with VCAM1 and GFAP antibodies and found that CCN1 was expressed in the surrounding ependymal cells, but not in VCAM1+ B1 cells located in the center of the pinwheel organization (Fig 1C). We traced the GFAP+ fibers of VCAM1+ B1 cells from the ventricular surface deep into the V‐SVZ in a series of confocal sections and found no CCN1 immunostaining signal in the cell bodies or processes of B1 NSCs (Fig 1D). We confirmed that CCN1 was only expressed in ependymal cells, but not in NSCs, oligodendrocytes, or neuroblasts based on the single‐cell RNA‐sequencing dataset from the adult V‐SVZ (Luo et al, 2015) (Fig EV1B). Additionally, we did not detect CCN1 in blood vessels (Fig EV1C). Thus, CCN1 expression is restricted to ependymal cells in the adult V‐SVZ.

Figure 1. CCN1 is specifically expressed in ependymal cells in the adult V‐SVZ.

Figure 1

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  1. Immunostaining for CCN1 (green) in the adult brain coronal section. Scale bar: 50 μm. The insert refers to the same image with DAPI staining to show cell nuclei (blue). LV, lateral ventricle.
  2. Adult V‐SVZ whole‐mounts stained for CCN1 (white) and S100β (green). Scale bar: 10 μm.
  3. Adult V‐SVZ whole‐mounts stained for CCN1 (white), GFAP (green), and VCAM1 (magenta). Arrowhead points to a VCAM1+ B1 cell surrounded by CCN1‐expressing ependymal cells. Scale bar: 10 μm.
  4. Confocal images tracing a VCAM1+ GFAP+ cell reveal CCN1 expression is absent in B1 cells. Yellow arrowheads point to the apical side of a B1 cell in the center of a NSC unit. White arrowheads point to the process of the B1 cell. Scale bar: 10 μm.

Figure EV1. Expression of CCN1 in the adult V‐SVZ.

Figure EV1

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  1. Whole‐mount staining for CCN1 (white) and β‐catenin (green) in the adult V‐SVZ. Scale bar: 10 μm.
  2. FPKM of Ccn1 mRNA in adult V‐SVZ cells (Luo et al, 2015). n = 10 (ependymal cells), 2 (neural stem cells), 9 (oligodendrocytes), and 11 (neuroblasts). Data are represented as mean ± SEM. *P = 0.0159, **P = 0.0045, ****P < 0.0001, n.s. not significant, one‐way ANOVA followed by Fisher's post hoc test.
  3. V‐SVZ whole‐mount staining for CCN1 (white) and CD31 (red), showing no CCN1 expression in the blood vessel. Scale bar: 10 μm.

Source data are available online for this figure.

CCN1 negatively regulates niche capacity and B1 cell number in the adult V‐SVZ

In the adult V‐SVZ, it has been shown that ependymal cells but not the B1 cells express the transcription factor FOXJ1 (Jacquet et al, 2009; Shah et al, 2018). To confirm the specificity of the Foxj1 CreERT2 mouse line (Muthusamy et al, 2014), we treated adult Foxj1 CreERT2 ; Rosa mT/mG mice with tamoxifen (TAM) for 5 consecutive days, which resulted in specific recombination in the ependymal layer, with 40% of the ependymal cells showing GFP expression (Fig EV2A).

Figure EV2. VCAM1 expression marks quiescent NSCs and NSC units.

Figure EV2

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  1. Specific labeling of ependymal cells in Foxj1 CreERT2; Rosa mTmG mice 1 week after TAM treatment. Scale bar: 20 μm.
  2. Western blotting showing reduced CCN1 protein level in the V‐SVZ of Ccn1cKO mice. n = 3, *P = 0.0123.
  3. Whole‐mount staining for γ‐tubulin (green), β‐catenin (white), and acetylated‐tubulin (magenta). Scale bar: 10 μm.
  4. Whole‐mount staining for γ‐tubulin (cyan), GFAP (red), and VCAM1 (white). Arrowheads point to VCAM1+ B1 cells in a NSC unit. Scale bar: 10 μm.
  5. Confocal images of adult V‐SVZ whole‐mounts showing scattered distribution of VCAM1+ B1 cells (arrowheads). Scale bar: 50 μm.
  6. Proportions of B1 cells that express VCAM1. n = 4, n.s. not significant.
  7. Whole‐mount staining for γ‐tubulin (cyan), β‐catenin (red), and VCAM1 (white). Arrowheads point to VCAM1+ NSC units. Scale bar: 10 μm.
  8. Average numbers of B1 cells in a NSC units. n = 4, **P = 0.0021.
Data information: In (B, F, and H), data are presented as mean ± SEM. Paired Student's t‐test (B). Unpaired Student's t‐test (F, H).Source data are available online for this figure.

To investigate the function of ependyma‐derived CCN1 in the adult V‐SVZ, we crossed Foxj1 CreERT2 mice with Ccn1 flox/flox mice (Kim et al, 2013) to specifically delete Ccn1 in ependymal cells. We treated both Foxj1 CreERT2; Ccn1 flox/flox (Ccn1cKO) and Ccn1 flox/flox (Ctrl) mice with TAM for 5 days at 8 weeks of age. We found that ependymal cells that had lost CCN1 expression distributed randomly in the ventricular surface, forming a mosaic pattern in the V‐SVZ 1 week post‐induction (Fig 2A). CCN1 protein level was notably reduced in the V‐SVZ of Ccn1cKO mice 1 week after TAM treatment (Fig EV2B). Loss of CCN1 did not lead to noticeable abnormalities in the shape and surface morphology of the V‐SVZ whole‐mounts, as revealed by reconstructed whole view of the lateral wall of the lateral ventricle (Fig 2B).

Figure 2. CCN1 restricts NSC pool size and niche capacity in the adult V‐SVZ.

Figure 2

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  1. V‐SVZ whole‐mount staining showing reduced CCN1 expression in Ccn1cKO mice 1 week after TAM treatment. Scale bar: 10 μm.
  2. V‐SVZ whole‐mounts stained for γ‐tubulin (cyan) and GFAP (red). Arrowheads point to B1 cells in the pinwheel center. Scale bars: 500 μm (left), 50 μm (middle), and 10 μm (right).
  3. Densities (cells/mm2) of B1 cells in the adult V‐SVZ. n = 4, ***P = 0.0005.
  4. V‐SVZ whole‐mounts stained for γ‐tubulin (cyan), GFAP (red), and VCAM1 (white). Arrowheads point to VCAM1+ B1 cells in NSC units. Scale bars: 50 μm (left) and 10 μm (right).
  5. Densities (cells/mm2) of VCAM1+ B1 cells in the adult V‐SVZ. n = 4, **P = 0.002.
  6. Densities (units/mm2) of NSC units in the adult V‐SVZ. n = 4, **P = 0.0022.
  7. Distribution of NSC units with different numbers of B1 cells. n = 4, ****P < 0.0001 (1 B1 cell/unit), **P = 0.0025 (2 B1 cells/unit), **P = 0.0063 (3 B1 cells/unit), *P = 0.0385 (4 B1 cells/unit).
Data information: In (C, E, F, and G), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

Remarkably, staining for GFAP in the whole‐mounts revealed significantly more GFAP+ cells in the V‐SVZ of Ccn1cKO mice compared with Ctrl mice (Fig 2B). We focused on the GFAP+ B1 cells, which bear a single primary cilium and are intermingled with multi‐ciliated ependymal cells. From the en face view of the V‐SVZ whole‐mounts, we found it difficult to discern the single cilium of B1 cells among tufts of ependymal motile cilia as revealed by acetyl‐tubulin staining (Fig EV2C), but γ‐tubulin staining, which labels the basal bodies of cilia, facilitated the identification and quantification of the ventricle‐contacting apical B1 cells in combination with GFAP staining (Fig 2B). We found that the number of GFAP+ cells with single γ‐tubulin+ basal body (B1 cells) was dramatically increased in the adult Ccn1cKO V‐SVZ compared with control (261.7 ± 45.5/mm2 in Ctrl; 871.7 ± 78.6/mm2 in Ccn1cKO) (Fig 2C).

To assess the impact on qNSCs, we stained whole‐mounts for VCAM1, which is expressed in the apical processes of qNSC (Fig EV2D). VCAM1+ qNSCs accounted for 63.69 ± 8.26% of total B1 cells in the V‐SVZ of control mice (Fig EV2E and F). There was a 3.5‐fold increase in the number of VCAM1+ B1 cells in CCN1‐deficient mice (168.3 ± 34.8/mm2 in Ctrl; 606.7 ± 76.6/mm2 in Ccn1cKO) (Fig 2D and E), while the percentage of VCAM1+ B1 cells was similar between Ccn1cKO and Ctrl mice (Fig EV2F). An overview of the VCAM1 staining signals revealed that VCAM1+ clusters were evenly spaced in the V‐SVZ whole‐mounts (Fig EV2E). Together with β‐catenin and γ‐tubulin staining, it is evident that VCAM1+ cell‐centered pinwheel structures dispersed over the ventricular surface in a unitary pattern, which we designated as NSC units (Fig EV2G). We found the density of NSC units was considerably higher in Ccn1cKO mice compared with controls (148.3 ± 30.5/mm2 in Ctrl; 396.7 ± 38.0/mm2 in Ccn1cKO) (Fig 2F). Furthermore, the average number of B1 cells within each unit was also notably increased in Ccn1cKO mice (Fig EV2H). While most of the units contained a single B1 cell in Ctrl mice, loss of CCN1 enabled NSC units to hold up to four B1 cells (Fig 2G). Thus, the level of CCN1 protein regulates the niche capacity for B1 NSCs in the adult V‐SVZ.

Enlarged niche capacity persists in the aging V‐SVZ of Ccn1cKO mice

To assess the long‐term effect of Ccn1 deletion on V‐SVZ niche capacity and NSC maintenance, we analyzed V‐SVZ whole‐mounts 16 months after TAM administration (Fig 3A). Remarkably, there were still more B1 cells and VCAM1+ B1 cells in the V‐SVZ of aged Ccn1cKO mice compared with aged controls (Fig 3A–C). The density of NSC units remained higher in aged Ccn1cKO mice (33.3 ± 1.7/mm2 in Ctrl; 103.3 ± 14.8/mm2 in Ccn1cKO) (Fig 3D), but the number of B1 cells contained in each unit dropped to levels indistinguishable from controls (Fig 3E). Interestingly, the density of VCAM1+ B1 cells was 3.6‐fold higher in aged Ccn1cKO mice compared with aged controls (33.3 ± 1.7/mm2 in Ctrl; 118.3 ± 14.2/mm2 in Ccn1cKO) (Fig 3B), which was similar to the fold change in young adult mice (Fig 2E). Thus, loss of CCN1 did not lead to unlimited increase in niche capacity, but the initially expanded niche and NSC pool were maintained in the V‐SVZ during aging.

Figure 3. The enlarged V‐SVZ niche persists till aging in Ccn1cKO mice.

Figure 3

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  1. Densities (cells/mm2) of B1 cells in the aged V‐SVZ. n = 3, **P = 0.0099.
  2. Densities (cells/mm2) of VCAM1+ B1 cells in the aged V‐SVZ. n = 3, **P = 0.0041.
  3. V‐SVZ whole‐mounts stained for γ‐tubulin (cyan), GFAP (red), and VCAM1 (white) in aged mice. Arrowheads point to VCAM1+ B1 cells in NSC units. Scale bars: 50 μm (left) and 10 μm (right).
  4. Densities (units/mm2) of NSC units in the aged V‐SVZ. n = 3, **P = 0.0093.
  5. Average numbers of B1 cells in a NSC unit. n = 3, n.s. not significant.
Data information: In (A, B, D, and E), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

Loss of CCN1 transiently expanded NSC pool at the expense of neurogenesis

To determine how the NSC pool was expanded in CCN1‐deficient V‐SVZ niche, we assessed NSC proliferation and differentiation at an earlier time point when B1 cell number had not reached the peak. Two days after TAM administration, we found a significant increase in B1 cell proliferation, with a concomitant reduction in the number of ASCL1‐expressing cells (C cells) in the Ccn1cKO V‐SVZ compared with controls (Fig 4A and B). We labeled newborn neurons with one injection of BrdU 2 days after TAM and analyzed the OB 4 weeks later. Consistently, loss of CCN1 reduced the number of BrdU‐labeled neurons in the OB granular layer (Fig 4C). To determine whether this was caused by increased cell death, we quantified apoptotic cells in the V‐SVZ of Ccn1cKO and Ctrl mice. There was no significant difference in the number of Caspase 3+ cells, suggesting that cell death is not a major reason for the decrease in neurogenesis (Fig EV3A). Thus, NSCs respond rapidly to CCN1 loss by increasing proliferation and reducing differentiation, which ultimately leads to NSC accumulation in the adult V‐SVZ.

Figure 4. Loss of CCN1 transiently enhanced NSC proliferation and reduced differentiation.

Figure 4

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  1. Densities (cells/mm2) of GFAP+ Ki67+ cells in V‐SVZ whole‐mounts 2 days after TAM treatment. n = 4, *P = 0.0141. V‐SVZ whole‐mounts stained for GFAP (red) and Ki67 (cyan). Arrowheads point to dividing GFAP+ cells. Scale bar: 10 μm.
  2. Numbers of ASCL1+ cells in V‐SVZ coronal sections 2 days after TAM treatment. n = 3, **P = 0.0087. V‐SVZ coronal sections stained for ASCL1 (red) and nuclei (blue). Scale bar: 50 μm.
  3. Densities (cells/mm2) of BrdU+ cells in the OB granule cell layer (GCL) 2 days after TAM treatment. n = 4 (Ctrl) and 3 (Ccn1cKO), **P = 0.0086. OB sections stained for BrdU (yellow) and nuclei (blue). Scale bar: 50 μm.
  4. Densities (cells/mm2) of GFAP+ Ki67+ cells in V‐SVZ whole‐mounts 4 weeks after TAM treatment. Arrowheads point to dividing GFAP+ cells. n = 4, n.s. not significant. Scale bar: 10 μm.
  5. Numbers of ASCL1+ cells in V‐SVZ coronal sections 4 weeks after TAM treatment. n = 3, n.s. not significant. Scale bar: 20 μm.
  6. Densities (cells/mm2) of BrdU+ cells in the OB GCL 4 weeks after TAM treatment. n = 4, n.s. not significant. Scale bar: 50 μm.
  7. Densities (cells/mm2) of GFAP+ Ki67+ cells in V‐SVZ whole‐mounts 16 months after TAM treatment. Arrowheads point to dividing GFAP+ cells. n = 4, n.s. not significant. Scale bar: 10 μm.
  8. Densities (cells/mm2) of BrdU+ cells in the OB GCL 16 months after TAM treatment. n = 4, n.s. not significant. Scale bar: 50 μm.
Data information: In (A‐H), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

Figure EV3. NSC expansion is restricted to a short period of time in CCN1‐deficient mice.

Figure EV3

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  1. Numbers of apoptotic cells in V‐SVZ coronal sections. n = 4, n.s. not significant. Confocal image shows apoptotic cells expressing cleaved caspase‐3 (red). Scale bar: 10 μm.
  2. Relative numbers of DCX+ neuroblasts in V‐SVZ coronal sections quantified at 2 days, 1 week, and 4 weeks after TAM treatment. n = 3 (2d, Ccn1cKO‐1w) and 4 (Ctrl‐1w, 1 m), *P = 0.0216, n.s. not significant.
Data information: In (A and B), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

Interestingly, NSC proliferation, production of C cells, and OB neurogenesis in CCN1‐deficient mice returned to a level similar to control mice 4 weeks after TAM treatment (Fig 4D–F). We also quantified DCX‐expressing neuroblasts in the V‐SVZ at different time points after TAM induction. Consistently, the number of DCX+ cells in Ccn1cKO mice was decreased 2 days after TAM and reverted to a level similar to that in Ctrl mice 1 week later (Fig EV3B). By 16 months after TAM treatment, neither NSC proliferation nor OB neurogenesis showed noticeable difference between Ccn1cKO and Ctrl mice (Fig 4G and H), suggesting that transient activation of NSCs in the early stage did not exhaust the stem cell pool and had no impact on the tissue homeostasis under normal conditions in the aged CCN1‐deficient V‐SVZ.

Loss of CCN1 increased functional quiescent NSCs

To determine whether the increased B1 cells are bona fide stem cells defined by function, we performed label‐retention and regeneration assays. Adult quiescent NSCs can be identified by BrdU label‐retention assays on the premise that these cells divide rarely (Morshead et al, 1998; Doetsch et al, 1999a). We performed BrdU‐labeling 2 days after TAM treatment by ten BrdU injections at an interval of 12 h (Fig 5A). After a 4‐week chase period, brain sections were double‐stained for both GFAP and BrdU to reveal label‐retaining qNSCs. We found significantly more BrdU‐retaining GFAP+ cells in Ccn1cKO mice compared with controls, indicating that the increased B1 cells in CCN1‐deficient mice returned to quiescence after initial activation and expansion (Fig 5A). Very few of these BrdU‐retaining cells were dividing in either Ctrl or Ccn1cKO mice (GFAP+ BrdU+ Ki67+/GFAP+ BrdU+ Ctrl: 1/130; Ccn1cKO: 0/194), indicating they have not been activated by this time point.

Figure 5. Loss of CCN1 increased quiescent NSCs in the V‐SVZ.

Figure 5

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  1. Numbers of GFAP+ BrdU+ label‐retaining NSCs in the V‐SVZ. Arrowheads point to label‐retaining NSCs. n = 4, *P = 0.039. V‐SVZ coronal sections stained for GFAP (cyan) and BrdU (red). Scale bar: 10 μm.
  2. Densities (cells/mm2) of GFAP+ Ki67+ cells in V‐SVZ whole‐mounts 24 h after TMZ treatment. Arrowheads point to dividing NSCs under regenerative condition. n = 6, *P = 0.0173. Scale bar: 10 μm.
  3. Densities (cells/mm2) of DCX+ neuroblasts in V‐SVZ whole‐mounts 3 days after TMZ treatment. n = 3, *P = 0.0179. V‐SVZ whole‐mounts stained for DCX. Scale bar: 500 μm.
  4. Densities (cells/mm2) of BrdU+ cells in the OB GCL labeled 24 h after TMZ treatment. n = 5 (Ctrl) and 6 (Ccn1cKO), *P = 0.0484. OB sections stained for BrdU (yellow) and nuclei (blue). Scale bar: 50 μm.
  5. Densities (cells/mm2) of GFAP+ Ki67+ cells in V‐SVZ whole‐mounts of aged mice 24 h after TMZ treatment. Arrowheads point to dividing NSCs under regenerative condition. n = 3, *P = 0.0321. Scale bar: 10 μm.
  6. Densities (cells/mm2) of DCX+ neuroblasts in V‐SVZ whole‐mounts of aged mice 3 days after TMZ treatment. n = 3, n.s. not significant. Scale bar: 500 μm.
  7. Densities (cells/mm2) of BrdU+ cells in the OB GCL of aged mice labeled 3 days after TMZ treatment. n = 4 (Ctrl) and 3 (Ccn1cKO), n.s. not significant. Scale bar: 50 μm.
Data information: In (A‐G), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

We labeled qNSCs 1 week before Ccn1 deletion and analyzed brain sections 4 weeks after TAM administration. There was no difference in the number of BrdU‐positive cells between Ccn1cKO and Ctrl mice (Fig EV4A), suggesting the pre‐existing qNSCs were not excessively consumed after the loss of CCN1.

Figure EV4. qNSCs are increased in CCN1‐deficient V‐SVZ.

Figure EV4

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  1. Numbers of BrdU‐retaining cells labeled before TAM treatment in V‐SVZ coronal sections. n = 6, n.s. not significant.
  2. Numbers of Ki67+ cells in V‐SVZ coronal sections 24 h after Ara‐C removal. n = 3, **P = 0.001. V‐SVZ coronal sections stained for Ki67 (red) and nuclei (blue). Scale bar: 50 μm.
  3. Densities (cells/mm2) of VCAM1+ B1 cells in the V‐SVZ 24 h after TMZ treatment. n = 6, ***P = 0.0004.
  4. Proportions of GFAP+ cells that express Ki67 24 h after TMZ treatment in young adult mice. n = 6, n.s. not significant.
  5. Proportions of GFAP+ cells that express Ki67 24 h after TMZ treatment in aged mice. n = 3, n.s. not significant.
Data information: In (A‐E), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

Quiescent NSCs are resistant to anti‐mitotic drug treatment and capable of regenerating the V‐SVZ after drug removal (Doetsch et al, 1999b; Mich et al, 2014). Four weeks after TAM administration, we infuse cytosine‐β‐D‐arabinofuranoside (Ara‐C) into the brain lateral ventricle for 6 days to kill all the dividing cells. Twenty‐four hours after pump removal, we detected a significantly higher number of dividing cells in the V‐SVZ of Ccn1cKO mice (Fig EV4B). Alternatively, mice received three daily doses of temozolomide (TMZ), another anti‐mitotic drug, starting 2 weeks after TAM administration (Fig 5B). 24 h after TMZ treatment, we found there were still more VCAM1+ B1 cells in Ccn1cKO mice (Fig EV4C). The number of proliferating GFAP+ cells was significantly higher in Ccn1cKO mice as compared with controls (Fig 5B). The proportion of GFAP+ cells that were Ki67+ was similar in both Ccn1cKO and Ctrl mice, indicating the increase in NSC proliferation was only related to increased size of the NSC pool, but not because GFAP+ B1 cells were proliferating more (Fig EV4D).

We then examined neuronal production under regenerative conditions by treating mice with TMZ 2 weeks after TAM (Fig 5C). Three days after the end of TMZ treatment, we found significantly more DCX+ immature neurons were generated in Ccn1cKO mice compared with controls (Fig 5C). We labeled newborn neurons by ten injections of BrdU 3 days after TMZ treatment and analyzed the OB 4 weeks later (Fig 5D). The density of BrdU‐labeled neurons in the OB granule layer was noticeably higher in CCN1‐deficient mice compared with controls (Fig 5D). Thus, loss of CCN1 increased quiescent NSCs capable of regenerating the adult V‐SVZ.

To determine whether the increased B1 cells were still functional NSCs in aged Ccn1cKO mice, we treated mice with TMZ 16 months after TAM administration (Fig 5E). Twenty‐four hours after TMZ treatment, we observed a higher number of dividing cells in Ccn1cKO mice compared with Ctrl (Fig 5E). The proportions of GFAP+ cells undergoing proliferation were similar between Ctrl and Ccn1cKO mice (Fig EV4E). However, 3 days after the end of TMZ treatment, we did not detect any noticeable difference in the generation of DCX+ immature neurons between Ccn1cKO and Ctrl mice (Fig 5F). We gave mice one injection of BrdU 3 days after TMZ treatment and analyzed newborn neurons in the OB 4 weeks later (Fig 5G). The densities of BrdU+ neurons in the OB granule layer were indistinguishable between Ccn1cKO and Ctrl mice (Fig 5G). Thus, although the increased B1 cells could be reactivated to proliferate, they had lost the neurogenic ability during aging in CCN1‐deficient mice.

EGFR signaling is required for NSC pool expansion in Ccn1cKO mice

Activation of EGFR signaling promotes NSC proliferation and reduces neuronal differentiation, leading to accumulation of NSC in the adult V‐SVZ (Doetsch et al, 2002; Gomez‐Gaviro et al, 2012), which is similar to the phenotypes observed in CCN1‐deficient mice. Also, CCN1 has been reported to suppress cell proliferation by inhibiting EGFR signaling (Chen et al, 2016). To determine whether deficiency of CCN1 increased NSCs through EGFR signaling, we examined EGFR expression in the V‐SVZ using FACS (Pastrana et al, 2009). hGFAP‐GFP transgenic mice were crossed with Ccn1cKO and Ctrl mice (Foxj1 CreERT2; Ccn1 flox/flox; hGFAP‐GFP called Ccn1cKO‐GFP mice and Ccn1 flox/flox; hGFAP‐GFP called Ctrl‐GFP mice, hereafter). Two days after TAM treatment, V‐SVZ cells freshly isolated from Ccn1cKO‐GFP and Ctrl‐GFP mice were stained with EGF‐AF555. Based on GFP and EGF‐AF555 fluorescent signals, V‐SVZ cells were separated into three populations: qNSCs (GFP+ EGFR−), aNSCs (GFP+ EGFR+), and type C cells (GFP− EGFR+) (Fig 6A). We found that deficiency of CCN1 did not alter the total number of V‐SVZ cells (Fig 6B). There were significantly more GFP+ cells in Ccn1cKO‐GFP mice (Fig 6C and D). We also found a 3.3‐fold increase in the frequency of qNSCs (GFP+ EGFR−) in Ccn1cKO‐GFP mice relative to controls (Fig 6C and D), which was consistent with the whole‐mount immunostaining results. Remarkably, the frequency of aNSCs (GFP+ EGFR+) was significantly higher in Ccn1cKO‐GFP mice as compared with controls (Fig 6C and D). On the contrary, the frequency of type C cells (GFP− EGFR+) was lower in the V‐SVZ of Ccn1cKO mice (Fig 6C and D). Together with the quantification data in Fig 4, these data indicate that deficiency of CCN1 increased EGFR‐expressing activated NSCs, promoting a switch of NSC fate from differentiation to self‐renewal.

Figure 6. CCN1 regulates NSC pool expansion through EGFR signaling.

Figure 6

Open in a new tab
  1. Representative FACS plots showing gating strategy for the analysis of cell types in the adult V‐SVZ.
  2. Total number of V‐SVZ cells isolated from one mouse. n = 3, n.s. not significant.
  3. Representative FACS plots showing aNSCs (GFP+ EGFR+), qNSCs (GFP+ EGFR−), and C cells (GFP− EGFR+) in the adult V‐SVZ.
  4. Percentage of each cell population in the adult V‐SVZ. n = 3, **P = 0.0014 (GFP+ EGFR−), ***P = 0.0002 (GFP+ EGFR+), *P = 0.0111 (GFP− EGFR+), ***P = 0.0006 (GFP+), n.s. not significant.
  5. Densities (cells/mm2) of VCAM1+ B1 cells in the adult V‐SVZ whole‐mounts. n = 4 (Ctrl + vehicle and Ccn1cKO + vehicle) and 3 (Ccn1cKO + erlotinib), **P = 0.0021 (Ccn1cKO + vehicle versus Ccn1cKO + erlotinib), **P = 0.0014 (Ccn1cKO + erlotinib versus Ctrl + vehicle), ****P < 0.0001.
  6. Densities (units/mm2) of NSC units in the adult V‐SVZ whole‐mounts. n = 4 (Ctrl + vehicle and Ccn1cKO + vehicle) and 3 (Ccn1cKO + erlotinib), *P = 0.017, **P = 0.0061, ***P = 0.0002.
  7. V‐SVZ whole‐mounts stained for γ‐tubulin (cyan), GFAP (red), and VCAM1 (white). Scale bar: 50 μm.
Data information: In (B, D, E, and F), data are presented as mean ± SEM. Unpaired Student's t‐test.Source data are available online for this figure.

To determine whether EGFR signaling is required for the expansion of B1 cell population in CCN1‐deficient niche, we gave mice five daily injections of erlotinib, a specific EGFR tyrosine kinase inhibitor, on the same day of TAM treatment with 12‐h intervals. Twenty‐four hours after drug treatment, we found that erlotinib reduced the densities of VCAM1+ B1 cells and NSC units in Ccn1cKO mice as compared with vehicle group and partially rescued the phenotype caused by CCN1 loss (Fig 6E–G). Thus, EGFR signaling is required for the increase of NSCs in CCN1‐deficient mice.

V‐SVZ niche expansion is caused by CCN1 loss, not other ependyma‐derived factors in Ccn1cKO mice

To determine whether the phenotypes observed in Ccn1cKO mice were caused by CCN1 loss directly or altered expression of other soluble proteins secreted by ependymal cells, we analyzed differentially expressed genes in ependymal cells of Ctrl and Ccn1cKO mice. We generated Foxj1 CreERT2; Rosa mT/mG; Ccn1 flox/flox (Ccn1cKO‐Rosa) and Foxj1 CreERT2; Rosa mT/mG; Ccn1 flox/+(Ctrl‐Posa) mice and treated the 8‐week‐old mice with TAM to delete Ccn1 and initiate GFP expression in ependymal cells. Two days after TAM treatment, we purified GFP+ ependymal cells from Ccn1cKO‐GFP and Ctrl‐GFP mice by FACS (Fig EV5A). Three samples from each genotype were processed for RNA‐sequencing and bioinformatical analyses. We identified 30 genes with more than 2‐fold change in expression level and P < 0.01 (Fig EV5B; Table EV1). Among the differentially expressed genes (DEGs), Ccn1 was the most downregulated in Ccn1cKO‐GFP mice, with the expression level reduced to 20% of Ctrl mice (Fig EV5B). GO analysis of downregulated genes in Ccn1cKO mice revealed marked enrichment in categories of protein transport and vesicle‐mediated transport (Fig EV5C). Upregulated genes did not reach significant enrichment in any GO terms. There was no significant difference in the expression of most secreted or membrane proteins, especially those known to regulate NSCs (Table EV2).

Figure EV5. NSC pool expansion is directly caused by CCN1 loss.

Figure EV5

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  1. Representative FACS plots showing gating strategy for the isolation of ependymal cells.
  2. Volcano plot of differentially expressed genes in ependymal cells. n = 3 for Ctrl and Ccn1cKO mice.
  3. Gene Ontology (GO) analyses of downregulated genes in Ccn1cKO ependymal cells.
  4. Densities (cells/mm2) of B1 cells in V‐SVZ whole‐mounts. n = 5 (Ctrl + saline and Ccn1cKO + CCN1) and 4 (Ccn1cKO + saline), **P = 0.0015, ***P = 0.0004, ****P < 0.0001. Data are presented as mean ± SEM. Unpaired Student's t‐test.

Source data are available online for this figure.

Thus, we hypothesized that loss of CCN1 is directly responsible for the increase in NSC population. Indeed, we found that exogenous CCN1 reduced the number of B1 cells and partially reversed the phenotype in Ccn1cKO mice after infusion of the recombinant CCN1 protein into the brain lateral ventricle during the 5‐day TAM injections compared with the vehicle (Fig EV5D).

CCN1 binds to integrin α6β1 in NSCs but the integrin signaling is not necessary for the effect on qNSC

NSCs in the adult V‐SVZ express integrin α6β1 (Shen et al, 2008; Kazanis et al, 2010) (Fig EV6A), which is a known binding target to mediate the functions of CCN1 in many other systems (Chen et al, 2000; Grzeszkiewicz et al, 2002; Lau, 2016). We found that CCN1 directly bound to integrin α6 and β1 subunits in V‐SVZ tissue lysates in a pull‐down assay using CCN1‐Fc protein (Fig EV6B). To determine the cell types that interacted with CCN1 in vivo, we performed the binding assay on V‐SVZ cells freshly isolated from hGFAP‐GFP transgenic mice. V‐SVZ cells were stained for CCN1‐Fc, integrin α6, and EGF‐AF555 ligand. qNSCs (GFP+ EGFR−) and aNSCs (GFP+ EGFR+) were separated depending on GFP and EGF‐AF555 fluorescence (Fig EV6C). We found that the majority of aNSCs (80.83 ± 0.23%) and a small percent of qNSCs (38.70 ± 2.07%) showed the binding of CCN1‐Fc and expression of integrin α6 (CD49f immunofluorescent signal) (Fig EV6D). Thus, CCN1 interacts with integrin α6β1 on NSCs in the adult V‐SVZ.

Figure EV6. Integrin α6β1 mediates the binding but not the function of CCN1 in the adult V‐SVZ.

Figure EV6

Open in a new tab
  1. Adult V‐SVZ whole‐mounts stained for γ‐tubulin (cyan), GFAP (red), and integrin α6 (white). Scale bar: 10 μm.
  2. Pull‐down experiment showing direct binding of CCN1 to integrin α6 and β1 subunits.
  3. qNSCs and aNSCs are separated based on GFP and EGFR expression.
  4. Percentages of cells that bind CCN1‐Fc and express integrin α6. n = 3.
  5. Densities (cells/mm2) of VCAM1+ B1 cells in the V‐SVZ after blocking integrin α6. n = 5, n.s. not significant.
  6. Densities (cells/mm2) of VCAM1+ B1 cells in the adult V‐SVZ after blocking integrin β1. n = 5, n.s. not significant.
Data information: In (D–F), data are presented as mean ± SEM. Unpaired Student's t‐test (E and F).Source data are available online for this figure.

To determine whether integrin α6β1 mediates the constraining effect of CCN1 on niche capacity and NSC pool, we infused blocking antibodies against integrin α6 and β1 into the brain lateral ventricle for 6 days, respectively. However, neither blocking antibodies altered the amount of VCAM1+ B1 cells in the V‐SVZ, indicating the effect of CCN1 was not mediated by integrin α6 or β1 subunits in the adult V‐SVZ (Fig EV6E and F).

Discussion

Adult somatic stem cells reside in specialized niches, where the abundance is tightly regulated and decreases gradually during aging to meet the homeostatic need of the tissue (Bouab et al, 2011; Conover & Shook, 2011b; Capilla‐Gonzalez et al, 2014; Obernier et al, 2018). Niche factors control the self‐renewal, proliferation, and differentiation of stem cells, playing an essential role in the regulation of adult stem cells. Here we show that the matricellular protein CCN1, which is selectively expressed in the ependymal cells, constrains the niche capacity, maintaining a balanced number of NSCs in the adult V‐SVZ.

In the adult V‐SVZ, the numbers of NSCs and NSC units decrease from P0 to P21 (Hu et al, 2017) and from young adult to aging mice (Shook et al, 2012). The abundance of NSCs contained in each unit also reduces during aging (Figs 2F and 4E). Most NSC‐activating factors promote NSC activation and consequently accelerate depletion. For example, infusion of EGF into the brain lateral ventricle increased the amount of B1 cells, which differentiated into astrocytes and lost neurogenic ability (Craig et al, 1996; Kuhn et al, 1997; Doetsch et al, 2002). Similarly, long‐term aberrant expression of cell‐intrinsic factors such as p57 rapidly increased NSCs, but eventually led to premature exhaustion of the stem cells at later stages (Kippin et al, 2005; Mira et al, 2010; Furutachi et al, 2013; Jones et al, 2015; Zhou et al, 2018). Interestingly, transient cell‐intrinsic expansion of adult NSCs by controlled over‐expression of Cdk4/cyclinD1 in NSCs leads to increased neurogenesis without depletion as a proportion of NSCs are able to return to quiescence (Bragado Alonso et al, 2019). We found that deficiency of CCN1 allows transient expansion of NSCs and renders the V‐SVZ niche capable of supporting more NSCs, resulting in more NSC units and NSCs per units. Our results present a niche regulatory strategy by which ependymal factor CCN1 controls the activation of NSCs and subsequent accommodation and maintenance of qNSCs in the apical niche surface of the V‐SVZ.

Upon CCN1 loss, the V‐SVZ niche was rapidly “enlarged” in terms of the number of B1 NSCs held within. Interestingly, the density of B1 NSCs reached a new homeostatic set point as the increased B1 cells entered a quiescent state, leading to preservation of qNSCs in the V‐SVZ until old age. Although the increased B1 cells in aged brain could be reactivated, they had lost neurogenic ability and could not contribute to more newborn neurons. This is reminiscent of the stem cell niche in Drosophila, where only daughter cells contacting hub cells preserve stem cell feature, ensuring that the primitive population is not beyond the nurturing limit (Xie & Spradling, 2000). Loss of multipotency is possibly due to inadequate amount of neurogenic factors in the niche to support a larger number of NSCs for a long time and/or the intrinsic loss of neurogenic potential in NSCs during aging. Thus, it would be interesting to see whether growth factors and morphogens could be used to restore their neurogenic ability.

The functions of CCN1 are mainly mediated by integrins, including integrin α6β1 (Lau, 2016). Integrin α6β1 is expressed in a variety of stem cell populations, including adult NSCs, and is crucial for stem cell maintenance and differentiation (Ramalho‐Santos et al, 2002; Krebsbach & Villa‐Diaz, 2017). During embryonic brain development, loss of integrin α6 results in abnormal cortical lamination and lack of integrin β1 affects NSC maintenance and neurogenesis (Georges‐Labouesse et al, 1998; Campos et al, 2004; Tan et al, 2016). In the adult V‐SVZ, integrin α6β1 is expressed in NSCs and mediates the adhesion to laminin‐rich blood vessels, blocking which promotes cell proliferation (Shen et al, 2008; Kazanis et al, 2010). However, the identity of the dividing cells is not clear. Here we found that infusion of blocking antibodies against integrin α6 or β1 had no effect on the number of ventricle‐contacting B1 NSCs in the adult V‐SVZ. Although CCN1 directly interacts with both α6 and β1 subunits in the adult V‐SVZ, they are not involved in the functions of CCN1 in regulating the NSC pool. Blockade of the integrin subunits did not alter the number of B1 cells, indicating other receptors might mediate the constraining effect of CCN1.

Signals from the niche determine the fate of stem cells, balancing between self‐renewal and differentiation. Once activated, neural stem cells start to express EGFR and enter a reversible state of quiescence (Pastrana et al, 2009; Codega et al, 2014). Activated NSCs also express transcription factor ASCL1, which is required for NSC proliferation and differentiation (Parras et al, 2004; Urban et al, 2016). Soon after Ccn1 deletion, EGFR‐expressing NSCs increased, and ASCL1+ cells decreased in the adult V‐SVZ, indicating CCN1‐deficient niche inhibited the lineage progression from EGFR‐expressing to ASCL1‐expressing cells. We did not find detectable interactions between CCN1 and EGFR or EGF in the CCN1‐Fc pull‐down assay, so it is likely that an indirect mechanism is involved.

The ependymal cells, intermingled with B1 NSCs along the ventricular surface, are a conceivable resource for niche factors that could affect NSC function. Surprisingly, we know so far less than a handful of ependyma‐derived factors that effectively influence the niche and NSC behavior. Our study has identified a novel ependyma‐specific niche factor and revealed that CCN1 imposes a constraining effect on V‐SVZ niche capacity and the size of NSC pool, implying the complexity of stem cell niche control.

Materials and Methods

Mice

All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Tsinghua University and under institutional assurances including AAALAC accreditation and PHS Animal Welfare Assurance (F16‐00228; A5061‐01). Mice were housed on a 12‐h light/dark cycle in standard cages with free access to water and food in the specific‐pathogen free facilities at Center of Biomedical Analysis in Tsinghua University. Both males and females at age 8–10 weeks were used. Ependyma‐specific Ccn1 knockout mice (Foxj1Cre ERT2; Ccn1 flox/flox) were generated by crossing Foxj1Cre ERT2 (generously provided by Ghashghaei, H. T.) (Muthusamy et al, 2014) with Ccn1 flox/flox (generously provided by Lau, L. F.) (Kim et al, 2013). ROSA mTmG (the Jackson Laboratory, Stock No. 007676) were maintained on a C57BL/6J background. hGFAP‐GFP mice (FVB/N‐Tg(GFAPGFP)14Mes/J, the Jackson Laboratory, Stock No. 003257) were maintained on a CD‐1 background. Both CD‐1 and C57BL/6J wild‐type mice were purchased from Vital River Laboratory Animal Technology Company (Beijing, China).

Drug administration

Tamoxifen (sigma, T5648) was dissolved in corn oil with 10% ethanol to prepare a 20 mg/ml solution and administered to both Ctrl and Ccn1cKO mice via intraperitoneal injections with 100 mg/kg for 5 consecutive days. BrdU (sigma, B5002) was dissolved in saline to get a 10 mg/ml solution, and mice were injected intraperitoneally with 100 mg/kg for indicated times as described in this article. TMZ (sigma, T2577) was dissolved in 25% DMSO/75% PBS to reach a final concentration of 10 mg/ml. Mice were injected intraperitoneally with 100 mg/kg/day for 3 consecutive days. Erlotinib (Selleck, S7786) was dissolved in 15% DMSO/85% captisol (Aladdin, C125030) at 15 mg/ml and administered via oral gavage with 150 mg/kg for 5 consecutive days.

Ara‐C, CCN1, and blocking antibody infusion

For V‐SVZ regeneration assay, osmotic pumps (ALZET model 1007D) filled with 2% Ara‐C (Sigma, C1768) were implanted onto the brain surface of adult mice following coordinates: anterior (A), 0; lateral (L), 0.8; and depth (D), 2.5 (relative to bregma and the surface of the brain) (Doetsch et al, 1999b). After the 6‐day infusion, the pumps were removed and mice were sacrificed 24 h later.

CCN1 (Peprotech, 120‐25) in saline (100 μg/ml) or saline alone was infused into the lateral ventricle with osmotic mini‐pumps at the same coordination for 6 days. Mice were treated with TAM for 5 consecutive days during protein infusion and sacrificed 24 h after the last TAM injection.

Blocking antibodies against integrin α6 (R&D Systems, MAB13501), integrin β1 (BD Pharmingen, 562219), isotype control rat IgG (R&D Systems, MAB006), and hamster IgG2 (BD Pharmingen, 553961) were dissolved in saline at 10 μg/ml and infused into the adult mice brain lateral ventricle for 6 days in osmotic pumps with the same coordination. Mice were sacrificed immediately after pump removal.

For all the experiments, contralateral hemispheres were used for further analysis.

Tissue processing, immunostaining, and imaging

For coronal section preparation, mice were deeply anesthetized with 1% pentobarbital sodium (i.p.) and transcardially perfused with saline, followed by 4% PFA. Brains were post‐fixed overnight at 4°C in 4% PFA and cryoprotected in 30% sucrose for 2 days at 4°C. Fixed tissues were embedded in OCT and cut into 30‐μm‐thick coronal sections. For fresh‐frozen section preparation, brains were extracted and immediately immersed in OCT. The mold containing the tissue was then fixed in a larger container, with the bottom contacting liquid nitrogen. When OCT was almost frozen, it was transferred to a −80°C refrigerator for further processing. For BrdU staining, sections were incubated in 2N HCL at 37°C for 30 min, followed by 0.1 M boric acid for 10 min at room temperature. Sections were blocked with 3% BSA containing 0.5% triton X‐100 and stained with primary antibodies overnight at 4°C followed by secondary antibodies for 2 h at room temperature. For whole‐mount staining, brains were extracted and the walls of the lateral ventricles were immediately dissected out (Mirzadeh et al, 2010). Whole‐mounts were fixed in 4% PFA overnight at 4°C. After washing in PBS, whole‐mounts were incubated in blocking solution (3% BSA with 2% triton X‐100) for 3 h at room temperature, and followed by primary and secondary antibodies for 24 h at 4 °C, respectively. After staining, the ventricular walls were dissected from underlying parenchyma and mounted. Primary antibodies used in this study are as follows: anti‐γ‐tubulin (Sigma, T5192), anti‐CCN1 (Abcam, ab24448), anti‐CCN1 (Santa Crus, sc‐8561), anti‐BrdU (Abcam, ab6326), anti‐Doublecortin (Santa Cruz, sc‐8066), anti‐GFAP (Sigma, G3893), anti‐GFAP (Millipore, AB5541), anti‐Ki67 (Lab Vision, RM‐9106‐S1), anti‐S100β (Sigma, AMAb91038), anti‐VCAM1 (BD Biosciences, 550547), anti‐integrin α6 (CST, 3750S), anti‐cleaved caspase 3 (CST, 9661S), and anti‐acetyl‐tubulin (Sigma, T6793). Secondary antibodies used are as follows: anti‐rabbit, anti‐mouse, anti‐goat, anti‐chicken, and anti‐rat antibodies conjugated with Alexa (AF488, AF568, or AF647) from Thermo Fisher Scientific. Images were acquired using Zeiss LSM780 Observer. Z2 and Zen software. Whole‐mount images were reconstructed from low power tiled confocal images acquired with the Plan‐Apochromat 20X/0.8 objective. For the analysis of B1 cells, high power images were taken from whole‐mounts with Plan‐Apochromat 63X/1.4 oil DIC objective. 10‐20 non‐overlapping fields across the entire whole‐mount were counted, and data were combined to determine the average intensity. Brain coronal sections were imaged with Plan‐Apochromat 40×/0.95 Corr objective. Images were analyzed using the Zeiss Zen and the Imaris software.

Pull‐down and Western blotting

For the analysis of CCN1 protein level, V‐SVZ whole‐mounts were dissected from Ccn1cKO and Ctrl mice 1 week after TAM treatment (Shen et al, 2008; Mirzadeh et al, 2010). Briefly, the walls of the lateral ventricles were dissected out from the caudal aspect of the telencephalon, septum, and hippocampus to expose the V‐SVZ whole‐mounts. V‐SVZ tissues covering the striatum were micro‐dissected from the corpus callosum to the ventral tip of the ventricle with minimal white matter and striatum parenchyma. The tissues containing ependymal cells and subependymal cells were lysed in 200 μl protein lysis buffer (1% NP‐40, 150 mM NaCl, 50 mM Tris‐Cl, pH 7.9) supplemented with protease inhibitor cocktail for 3 h at 4°C. Protein was extracted from the lysates by centrifuging at 12,000 g for 20 min at 4°C.

For pull‐down assay, V‐SVZ whole‐mounts were dissected from 8‐week‐old CD‐1 mice, and protein was extracted with the similar method mentioned above. 30 μg CCN1‐Fc (R&D systems, 4055‐CR) or Fc (R&D systems, 110‐HG) was incubated with Protein G dynabeads (Invitrogen, 10004D) in lysis buffer for 3 h at 4°C. After washing, beads were incubated with V‐SVZ protein extract overnight at 4°C, washed three times in lysis buffer, and eluted in SDS loading buffer.

For Western blotting, samples were separated on SDS–PAGE and transferred onto PVDF membrane. After blocking in 5% BSAT, membranes were incubated in primary antibody overnight at 4°C followed by secondary antibodies for 1 hr at room temperature. Primary antibodies used in this study are as follows: anti‐integrin α6 (CST, 3750S), anti‐integrin β1 (Millipore, MAB1997), and anti‐CCN1 (Santa Crus, sc‐8561). Signal was detected with corresponding HRP‐conjugated secondary antibodies and ECL immunoblotting detection system (Millipore).

FACS analysis

The V‐SVZs were dissected, minced, and digested in 0.05% trypsin‐EDTA with 20 μg/ml DNase 1 at 37°C for 3 min. Digested V‐SVZ pieces were gently triturated and washed three times in DMEM. The cell suspension was filtered through a 40 μm mesh to remove large tissue pieces.

For V‐SVZ cell lineage analysis, dissociated cells in 200 μl staining buffer were incubated with EGF‐AF555 (Thermo Fisher Scientific, E35350) (1:300) for 20 min at 4°C. After washing, cells were pelleted in 400 μl staining buffer containing 7AAD (1:200) and analyzed on LSRFortessa SORP.

For in vivo CCN1‐Fc binding assay, dissociated cells in 200 μl staining buffer were first incubated with CCN1‐Fc or Fc for 20 min at 4°C. After washing, cells were resuspended in 200 μl staining buffer and incubated with anti‐Human IgG‐AF647 (Jackson ImmunoResearch, 709‐605‐149), EGF‐AF555 and anti‐CD49f‐PE‐Cy7 (eBioscience, 25‐0495‐82), or anti‐Rat IgG‐PE‐Cy7 (eBioscience, 25‐4321‐82) for 20 min at 4°C. After washing, cells were pelleted in 400 μl staining buffer containing DAPI (1:5,000) and analyzed on LSRFortessa SORP.

Gates were set manually by using control samples, and data were analyzed with FlowJo V10 software.

Ependymal cell sorting

The V‐SVZs from Foxj1 CreERT2; Ccn1 flox/flox; ROSA mT/mG, Ccn1 flox/flox; ROSA mT/mG, and wild‐type C57BL/6J mice were dissected and digested in 0.05% trypsin‐EDTA with 20 μg/ml DNase 1 at 37°C for 1 min, which was enough to dissociate the ependymal cells without causing damage. Cell suspension was filtered through a 40 μm mesh and resuspended in DAPI‐containing buffer (1:5,000). FACS was performed on the BD INFLUX cell sorter using 10 psi pressure and 100 μm nozzle aperture. Gates were set manually by using control samples. About 1,000 ependymal cells from each mice were directly sorted into RLT lysis buffer.

Library preparation and sequencing

RNA‐Seq libraries were prepared from three biological replicates for each genotype. RNA from ependymal cells was extracted with the RNeasy Plus Micro Kit (QIAGEN, 74034). The whole transcriptome amplification products were generated using the Discover‐sc™ WTA Kit V2 (Vazyme, N711). A total amount of 1 ng qualified WTA cDNA products per sample was used as input material for library preparation. The sequencing libraries were generated using the TruePrep DNA Library Prep Kit V2 for Illumina® (Vazyme, TD503) following manufacturer's recommendations. Library concentration was preliminarily measured using Qubit® DNA Assay Kit in Qubit® 3.0, and the insert size was assessed using the Agilent Bioanalyzer 2100 system. The libraries were pooled at equal concentration following accurate quantification by StepOnePlus Real‐Time PCR system (ABI, USA). After cluster generation on cBot Cluster Generation System (Illumina), libraries were sequenced on Illumina HiSeq X Ten platform with 150 bp paired‐end module.

Quantification and statistical analysis

Statistical analysis was performed using two‐tailed Student's t‐test for two‐group comparison and one‐way ANOVA with Fisher's post hoc test for multiple comparisons. Results were analyzed using GraphPad Prism version 6 (GraphPad Software, Inc.). Data are represented as mean ± SEM. Significance is defined as n.s. P > 0.05, *P ≤ 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

Author contributions

QS conceived the idea and supervised the project. JW performed and interpreted histology, cytometry, surgeries, and mice breeding. W‐JT performed pull‐down and Western blot experiments. YL and XW performed RNA‐sequencing analysis. JLi conducted mice genotyping experiments. W‐JT, JZ, HP, HJW, and JLu helped with experiments and figure preparing. LFL and HTG provided mouse lines. HTG and XY discussed the data and assisted with manuscript preparation. JW and QS wrote the manuscript.

Conflict of interest

The authors declare that they have no conflict of interest.

Supporting information

Expanded View Figures PDF

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Table EV1

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Table EV2

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Source Data for Expanded View

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Review Process File

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Source Data for Figure 2

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Acknowledgements

We thank Yue Wang for technical assistance in osmotic pump infusion. We thank Core Facility of Center of Biomedical Analysis, Tsinghua University for assistance with Confocal Microscopy and Flow cytometry analysis. We are grateful for the service provided by Laboratory Animal Research Center at Tsinghua University. We also thank Xinjie Bao and Renzhi Wang for their help with discussion and funding support. This work was supported by the Roche‐Tsinghua collaborative grant, the Fundamental Research Funds for the Central Universities (22120190149), the National Key R&D Program of China (2018YFA0108600) and Ministry of Science and Technology (2019YFA0110102). H.T.G is supported by grants from the National Institutes of Health (R01NS098370 and R01NS089795).

The EMBO Journal (2020) 39: e101679

Data availability

Sequencing data have been deposited at GEO under accession number GSE137852 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE137852).

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Associated Data

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Supplementary Materials

Expanded View Figures PDF

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Table EV1

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Table EV2

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Source Data for Expanded View

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Review Process File

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Source Data for Figure 6

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Data Availability Statement

Sequencing data have been deposited at GEO under accession number GSE137852 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE137852).

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