Neural stem/progenitor cell (NSPC) proliferation and self‐renewal, as well as insult‐induced differentiation, decrease markedly with age. The molecular mechanisms responsible for these declines remain unclear. Here, we show that levels of NAD+ and nicotinamide phosphoribosyltransferase (Nampt), the rate‐limiting enzyme in mammalian NAD+ biosynthesis, decrease with age in the hippocampus. Ablation of Nampt in adult NSPCs reduced their pool and proliferation in vivo. The decrease in the NSPC pool during aging can be rescued by enhancing hippocampal NAD+ levels. Nampt is the main source of NSPC NAD+ levels and required for G1/S progression of the NSPC cell cycle. Nampt is also critical in oligodendrocytic lineage fate decisions through a mechanism mediated redundantly by Sirt1 and Sirt2. Ablation of Nampt in the adult NSPCs in vivo reduced NSPC‐mediated oligodendrogenesis upon insult. These phenotypes recapitulate defects in NSPCs during aging, giving rise to the possibility that Nampt‐mediated NAD+ biosynthesis is a mediator of age‐associated functional declines in NSPCs.
See also: C Wiley & J Campisi (June 2014)
Levels of NAD+ and the NAD+ biosynthetic enzyme Nampt decline in the hippocampus during aging. Nampt ablation in adult neural stem/progenitor cells reduces self‐renewal, proliferation, and oligodendrogenesis. Age‐associated decreases in the stem cell pool are reversed by the administration of nicotinamide mononucleotide, a key NAD+ intermediate.
Nampt is expressed in adult neural/progenitor stem cells and is the main source of their NAD+ levels.
Loss of Nampt reduces proliferation by stalling progression of the neural stem/progenitor cell cycle at G0/G1.
The NAD+‐dependent sirtuins Sirt1 and Sirt2 redundantly mediate neural stem/progenitor cell fate decisions into oligodendrocytes.
Loss of Nampt reduces neural stem/progenitor cell‐derived oligodendrocyte regeneration after cuprizone‐induced demyelination.
Aging manifests as a physiological breakdown of robustness, leading to a significant functional decline, increased vulnerability to environmental insults, and many different diseases in a variety of tissues and organs. Cognitive decline is one such example, and cognitive impairment, even without dementia, occurs in 22% of people over age 71 in the United States (Plassman et al, 2008). With this prevalence, efforts to understand the mechanisms underlying age‐associated cognitive decline and to develop therapeutic strategies to prevent it have become particularly poignant. With these goals in mind, the regulation of adult neurogenesis has been an intense focus of research. Throughout the lives of all mammals, including humans, two distinct populations of neural stem cells (NSCs) are maintained in the brain: one in the subgranular zone (SGZ) of the dentate gyrus (DG) and the other in the subventricular zone (SVZ) bordering the lateral ventricles. NSCs have the ability to self‐renew and to differentiate into transient amplifying progenitors (collectively referred to as neural stem/progenitor cells or NSPCs), which in turn undergo finite, lineage‐restricted cell divisions to produce neurons, oligodendrocytes, or astrocytes (Deng et al, 2010; Artegiani & Calegari, 2012; Jadasz et al, 2012).
Aging is one of the strongest negative regulators of adult NSPC proliferation (Artegiani & Calegari, 2012). In mice, the number of proliferating NSPCs in the SGZ declines exponentially in the first 9 months of life (Ben Abdallah et al, 2010), and they are almost completely absent by 24 months of age (Jin et al, 2003; Lugert et al, 2010; Encinas et al, 2011). Aging is also associated with impaired oligodendrocyte differentiation and remyelination in response to insult (Sim et al, 2002; Franklin & Ffrench‐Constant, 2008). Given the potential of NSPCs to contribute to cognitive function and their ability to proliferate and differentiate into the major cell types of the brain (Deng et al, 2010; Artegiani & Calegari, 2012; Jadasz et al, 2012), it is of great importance to understand key signaling pathways regulating the NSPC decisions of proliferation versus quiescence and self‐renewal versus terminal differentiation. NSPCs can be reactivated in the aged brain (Decker et al, 2002; Jin et al, 2003; Lugert et al, 2010). Therefore, elucidation of the key signaling pathways that are involved in age‐associated decline in NSPC functionality will provide critical insight into preventing age‐associated cognitive decline.
We have previously found that aging significantly reduces levels of the essential cofactor nicotinamide adenine dinucleotide (NAD+) in multiple peripheral tissues (Yoshino et al, 2011). This age‐associated decrease in NAD+ levels is due to a decline in protein levels of nicotinamide phosphoribosyltransferase (Nampt), the rate‐limiting enzyme in the biosynthetic pathway of NAD+ from nicotinamide (Fig 1A) (Revollo et al, 2004; Yoshino et al, 2011). Nampt converts nicotinamide, a major precursor in mammalian NAD+ biosynthesis, and 5′‐phosphoribosyl‐1‐pyrophosphate to nicotinamide mononucleotide (NMN), a key NAD+ intermediate. NMN is then converted to NAD+ by nicotinamide/nicotinic acid mononucleotide adenylyltransferase (Nampt) (Revollo et al, 2004, 2007). We and others have previously reported that the expression of Nampt in the brain is extremely low compared to peripheral tissues (Revollo et al, 2007; Friebe et al, 2011). However, Nampt has uniquely strong expression in the hippocampus (Zhang et al, 2010; Wang et al, 2011a). Because recent studies show that the energetic demands of stem cell proliferation and lineage specification require distinct metabolic programs (Folmes et al, 2012), we hypothesized that NSPCs would be particularly sensitive to changes in NAD+ levels and accordingly alter their proliferation, self‐renewal, and differentiation.
Here, we show that hippocampal NAD+ and Nampt levels decrease with age. Nampt mediates NSPC NAD+ biosynthesis. Pharmacological inhibition and genetic ablation of Nampt‐mediated NAD+ biosynthesis in NSPCs impair NSPC proliferation, self‐renewal, and formation of oligodendrocytes in vivo and in vitro. Furthermore, augmentation of NAD+ levels during aging with NMN administration maintains the NSPC pool. Thus, enhancing NAD+ levels in NSPCs may be an effective intervention to preserve the endogenous NSPC population to repair the aged, diseased, or damaged brain.
Hippocampal NAD+ levels and Nampt expression decline with age
Since aging significantly reduces levels of Nampt and NAD+ in multiple peripheral tissues (Yoshino et al, 2011), we hypothesized that aging also reduces Nampt‐mediated NAD+ biosynthesis in the brain, particularly in the hippocampus, affecting the function of NSPCs. To address this hypothesis, we first measured NAD+ levels in hippocampi isolated from 1‐, 3‐ to 4‐, 6‐, and 10‐ to 12‐month‐old C57Bl6 mice. NAD+ levels gradually decreased with age, reaching 63% in 10‐ to 12‐month‐old mice compared to that of 1‐month‐old mice (Fig 1B). Consistent with this finding, quantifying Nampt immunoreactivity in the SGZ of the DG by both a thresholded level of Nampt intensity and a count of the number of thresholded Nampt+ cells demonstrated that 18‐month‐old mice exhibit 52–66% of the Nampt immunoreactivity present in 6‐month‐old mice (Fig 1C–E). These results suggest that Nampt‐mediated NAD+ biosynthesis in the hippocampus declines with age at a time course similar to that of NSPC proliferation (Ben Abdallah et al, 2010).
Nampt is expressed in a subpopulation of SGZ NSPCs
Previously, it was reported that Nampt is predominantly expressed in hippocampal neurons, but not in stellate astrocytes (Zhang et al, 2010; Wang et al, 2011a). Consistent with this finding, immunohistochemistry for Nampt and cell type‐specific markers revealed that almost all NeuN+ neurons in the granule layer of the DG expressed Nampt, while almost no S100β+ glial cells did (Supplementary Fig S1A–E). However, we also noticed that many intensely Nampt immunoreactive cells along the SGZ of the DG did not express NeuN (Supplementary Fig S1B and E). Since NSPCs engage in the energetically costly tasks of proliferation and differentiation (Folmes et al, 2012), we performed co‐immunohistochemistry for NSPC markers (Sox2+, radial Gfap+) and found that a majority of NSPCs expressed Nampt (Fig 2A–D). To assess in vivo colocalization between Nampt and Nestin, we crossed mice expressing Cre recombinase under the Nestin promoter (Nestin‐CreERT2) (Lagace et al, 2007) to a GFP reporter mouse strain that expresses a loxP‐flanked STOP cassette that prevents transcription of the downstream enhanced GFP (see the Materials and Methods section), generating iNSPC‐GFP mice. Nampt also colocalized with GFP driven by the Nestin promoter (NestinGFP, Fig 2C and D) (Lagace et al, 2007). Quantification of these observations revealed that along the SGZ, 32% of Sox2+ cells, 55% of radial Gfap+ cells, and 78% of NestinGFP+ cells expressed Nampt (Fig 2D, Supplementary Fig S1F). Additionally, Ki67+ and Olig2+ cells along the SGZ also expressed Nampt (Supplementary Fig S1G–I). To confirm that Nampt is highly expressed in NSPCs, we cultured NSPCs from the hippocampi of postnatal pups as neurospheres. Neurospheres showed 22 or 32% higher expression levels of Nampt than did whole hippocampal extracts taken from postnatal (P12) or adult mice (2.5–4.5 months), respectively (Fig 2E), indicating that NSPCs have higher expression levels of Nampt compared to other hippocampal cell types. Together, these results suggest that Nampt is expressed in a large subpopulation of NSPCs.
Having found that Nampt expression is present in both neurons and NSPCs, we next asked which cell populations lose Nampt expression with age. To address this question, we thresholded Nampt immunoreactivity and assessed the thresholded Nampt+ cells for colocalization with the neuronal marker NeuN and the NSPC marker Sox2. With age, the percentage of intensely Nampt immunoreactive cells that colocalized with NeuN increased slightly, whereas the percentage of intensely Nampt immunoreactive cells that colocalized with Sox2 decreased from 21 to 4% (Fig 2F). Similarly, in the SGZ, the percentage of NeuN+ that expressed Nampt increased with age, while the percentage of Sox2+ cells that expressed Nampt decreased (Supplementary Fig S1E and F). Thus, at least part of the decrease in Nampt expression in the SGZ with age is due to the loss of expression from Sox2+ NSPCs.
Adult NSPC‐specific deletion of Nampt impairs NSPC self‐renewal in vivo
Having seen enrichment of Nampt in NSPCs, we next asked whether inactivating Nampt specifically in adult NSPCs could recapitulate age‐associated phenotypic changes in NSPC functionality in vivo. To address this question, we generated adult NSPC‐specific inducible Nampt‐knockout mice by crossing Namptflox/flox mice (Rongvaux et al, 2008) with Nestin‐CreERT2 mice (iNSPC‐Nampt‐KO mice). To trace the progeny of adult NSPCs in which Nampt was inactivated and to confirm the specificity and magnitude of the deletion induced by tamoxifen, we also crossed iNSPC‐Nampt‐KO mice to the aforementioned iNSPC‐GFP mice. After tamoxifen injection, these mice expressed NestinGFP in the SGZ and SVZ but not in non‐neurogenic regions of the brain such as the corpus callosum or cortex (Supplementary Fig S2A and B). Immunohistochemistry and recombination PCR for NestinGFP confirmed that there was undetectable recombination present in vehicle‐injected mice (Supplementary Fig S2A and C). To verify that the NestinGFP+ population consisted of NSPCs, we costained for the NSPC markers Sox2 and Gfap. 61% of Sox2+ cells and 34% of radial Gfap+ cells coexpressed NestinGFP 7 days post‐tamoxifen (Supplementary Fig S2D). We also verified Nampt deletion efficiency by quantifying the percentage of NestinGFP+ cells that expressed Nampt 3 and 7 days post‐tamoxifen injection. At 3 days post‐tamoxifen injection, the percentage of NestinGFP+ cells that expressed Nampt in iNSPC‐Nampt‐KO mice was 40% less than littermate controls, and at 7 days post‐tamoxifen injection, the percentage of NestinGFP+ cells that expressed Nampt was reduced by 62% (Supplementary Fig S2E).
To assess the cumulative effect of loss of Nampt on NSPC proliferation, we deleted Nampt in iNSPC‐Nampt‐KO mice at 6 weeks of age with three rounds of five consecutive days of tamoxifen injections, separated by 6 weeks (Fig 3A). We then assessed control and iNSPC‐Nampt‐KO mice for the expression of lineage‐specific markers by immunohistochemistry (Fig 3B). In iNSPC‐Nampt‐KO mice, we found that the Nestin+ NSPC pool was significantly decreased by 49% in the DG (Fig 3C). Indeed, incorporation of BrdU and the population of proliferating cells [Ki67+ (von Bohlen und Halbach, 2011)] were also decreased by 22% (P = 0.019) and 35% (P = 0.064), respectively (Fig 3D and E). Consistent with this defect in the NSPC pool and proliferation, the pool of newborn neurons [doublecortin, Dcx+ (von Bohlen und Halbach, 2011)] was also significantly decreased by 26% (Fig 3F and G). In contrast, we did not observe any significant difference in the maturation of newborn neurons, as assessed by categorization of Dcx+ cells with no or horizontal projections as immature neurons and Dcx+ cells with vertical projections spanning the granule cell layer as mature neurons (Supplementary Fig S2F). We next assessed NSPC/daughter cell survival by immunostaining for activated caspase‐3. Only rare activated caspase‐3+ cells were observed in both neurogenic and non‐neurogenic regions of the brain (Supplementary Fig S2A and B), and these rare activated caspase‐3+ cells were never observed in GFP+ cells in iNSPC‐Nampt‐KO DG, providing evidence against a potential contribution of cell death to the observed effects.
To assess the acute effect of loss of Nampt on NSPC fate decisions, we induced deletion of Nampt at 6 weeks of age with four total tamoxifen injections followed by sacrifice 72 h after the first injection (Fig 3H). To facilitate assessment of differentiation, we labeled dividing cells by injecting the mice with BrdU concurrently with the first day of tamoxifen treatment. iNSPC‐Nampt‐KO mice displayed significantly reduced levels of colocalization of BrdU with radial Nestin+ cells (Fig 3I), suggesting decreased self‐renewal decisions. However, iNSPC‐Nampt‐KO mice exhibited normal levels of BrdU colocalization with neuronal (Dcx+), astrocytic (Gfap+, also marks NSPCs), and oligodendrocytic (Olig2+) markers, indicating that alterations in differentiated cell lineage decisions were undetectable under basal conditions. The lack of increase in colocalization of BrdU with cell type‐specific markers may imply that a larger percentage of BrdU+ cells have failed to differentiate in iNSPC‐Nampt‐KO mice. iNSPC‐Nampt‐KO NSPCs could have stalled during differentiation after losing Nestin expression.
While NSPC proliferation declines exponentially throughout life (Artegiani & Calegari, 2012), quiescent NSPCs can be reactivated in the aged murine hippocampus by multiple environmental stimuli (Decker et al, 2002; Jin et al, 2003; Lugert et al, 2010). If NSPCs quiesce due to reduced NAD+ levels, systemic administration of NMN may be able to correct age‐associated defects in NSPC functionality. Intraperitoneal injection of NMN (500 mg/kg body weight) increased hippocampal NAD+ levels 34–39% within 15 min, strongly suggesting that NMN can cross the blood–brain barrier (Supplementary Fig S2G). To see whether NMN supplementation can maintain NSPC proliferation and self‐renewal with age, we treated 6‐month‐old mice with NMN at the daily dose of 100 or 300 mg/kg body weight in their drinking water until 18 months of age. The number of Nestin+ cells along the SGZ was significantly lower in the 18‐month‐old control mice relative to 6‐month‐old mice, as previously reported (Encinas et al, 2011) (Fig 3J). Remarkably, mice treated with 300 mg/kg body weight NMN showed improved maintenance of the type 1 (radial Nestin+) population with age. However, the population of proliferating cells (Ki67+) remained similar to controls (Supplementary Fig S2H). While not statistically significant, the population of newborn neurons (Dcx+) tended to increase (Supplementary Fig S2I). The age‐related depletion of the NSPC pool is thought to be caused by an increase in terminal fate decisions relative to self‐renewal fate decisions (Encinas et al, 2011). Thus, it is possible that NMN administration maintains the NSPC pool by preventing the age‐associated increase in terminal fate decisions.
Inhibition of Nampt in NSPCs in vitro impairs NAD+ biosynthesis and proliferation
Having shown that NAD+ levels in the hippocampus and Nampt expression in the SGZ decreased with age, we next asked whether Nampt mediates NSPC‐specific NAD+ biosynthesis by using hippocampal neurospheres as the in vitro NSPC culture model. We treated neurospheres with a highly specific Nampt inhibitor, FK866, at a dosage and duration (10 nM, 48 h) that has little to no effect on cellular viability (Hasmann & Schemainda, 2003). Strikingly, FK866 reduced NAD+ levels in neurospheres to 4% of controls, a decrease completely rescued by concurrent NMN treatment ((Fig 4A, Supplementary Fig S3A), strongly suggesting that Nampt activity is the predominant source of NAD+ biosynthesis in NSPCs.
We next examined how inhibition of Nampt affects neurosphere proliferation. Consistent with the decreases in the NSPC pool and in NSPC proliferation in iNSPC‐Nampt‐KO mice, FK866 reduced NSPC number by 61% after 48 h, but not 24 h, of treatment (Fig 4B and C, Supplementary Fig S3B). To distinguish whether this decrease in cell number was due to an inhibition of proliferation or enhancement of death, we analyzed the protein levels of markers of proliferation, apoptosis, and autophagy. Expression of the proliferation markers Ki67 and PCNA decreased 87 and 43%, respectively (Supplementary Fig S3C–E), whereas levels of activated caspase‐3 became detectable and levels of the autophagy marker, glycosylated LC3B, were unchanged. Consistent with these observations, parametric analysis of gene set enrichment (PAGE) of a microarray performed on neurospheres treated with FK866 showed that out of the top 50 downregulated pathways, 13 of them were related to the cell cycle, while none of the top 50 upregulated pathways were involved in cell death (Fig 4D, Supplementary Fig S3F and G). Analysis of specific gene changes by qRT–PCR revealed that cyclins E and A, the two cyclins required for cellular progression from G1 to S, as well as their upstream transcriptional regulator E2F1 (Wong et al, 2011), were the primary cell cycle factors affected by this treatment (Fig 4E). These alterations in gene expression indicated that reducing Nampt activity stalls NSPCs at G0/G1. Supporting this notion, FACS analysis of neurospheres demonstrated that FK866 treatment increased the proportion of NSPCs in G0/G1 and decreased the proportion in S phase (Fig 4F).
Genetic ablation of Nampt in NSPCs in vitro impairs NAD+ biosynthesis, proliferation, and differentiation
To assess the effect of chronic Nampt ablation on NSPC functionality, we genetically ablated Nampt by infecting neurospheres from Namptflox/flox mice with Cre recombinase‐ or LacZ‐expressing (control) adenoviruses. Neurospheres infected with Cre recombinase (Nampt Ad‐Cre) at passage 1 exhibited a 94% reduction in Nampt mRNA expression 3 days post‐deletion, and the corresponding decreases in Nampt protein expression and NAD+ levels appeared 6 days post‐deletion (Supplementary Fig S4A–E). Eight days post‐deletion, NSPCs exhibited a 73% reduction in NAD+ levels that was rescued by concurrent NMN administration, further supporting the notion that Nampt activity is the predominant source of NSPCs NAD+ levels (Fig 5A).
Like FK866‐treated cultures, proliferating Nampt Ad‐Cre‐infected NSPCs displayed reduced cell number (Fig 5B). Remarkably, Nampt Ad‐Cre NSPCs were unable to increase their cell number between 24 and 144 h of culture. In contrast, Nampt AD‐LacZ‐infected cells were able to exponentially increase their cell number over 13‐fold in this time frame. Consistent with this finding, Nampt Ad‐Cre‐infected NSPCs also showed a 49% reduction in diameter relative to Nampt AD‐LacZ‐infected NSPCs, indicative of reduced proliferation (Fig 5C and D). Since NSPC self‐renewal decisions can also contribute to cell number, we assessed secondary neurosphere formation, an assay that quantifies the ability of neurosphere inhabitant cells to reformulate neurospheres upon dissociation. Nampt Ad‐Cre‐infected cells generated 63% fewer secondary neurospheres than did Nampt AD‐LacZ‐infected cells (Fig 5E). Nampt AD‐LacZ and Nampt Ad‐Cre NSPCs exhibited no difference in the percentages of TUNEL‐ or activated caspase‐3‐positive cells as well as no difference in activated caspase‐3 immunoreactivity as detected by immunoblotting, indicating that the observed phenotypes upon loss of Nampt are not primarily due to cell death (Supplementary Fig S4E and F). To see whether Nampt Ad‐Cre‐infected neurospheres can be reactivated to proliferate, we plated equal numbers of Nampt AD‐LacZ and Nampt Ad‐Cre cells after the second passage and cultured them in the presence or the absence of NMN. Remarkably, NMN treatment was able to fully reactivate the proliferative potential of Nampt Ad‐Cre cells (Fig 5F and G). Collectively, these results suggest that Nampt‐mediated NAD+ biosynthesis is critical for NSPCs to successfully progress through the cell cycle.
Whereas we did not observe a difference in NSPC fate decisions in the neurogenic environment of the SGZ in vivo, we detected a decrease in self‐renewal decisions. To see whether this was true in the absence of the influences of the SGZ niche, we differentiated dissociated neurospheres and assessed the proportion of resulting cell types by immunofluorescence after 6–7 days of differentiation induced by the removal of growth factors (Fig 5H, Supplementary Fig S4G). To our surprise, differentiated Nampt Ad‐Cre NSPCs exhibited a 90% reduction in oligodendrocytes (O4+, Fig 5I). In contrast, Nampt Ad‐Cre‐infected NSPCs exhibited no change in the generation of Gfap+ cells (Fig 5J). Genetic knockdown of Nampt also significantly but more mildly decreased the generation of neurons (43% decrease, β‐III‐tubulin+, Fig 5K). Thus, the decrease in oligodendrocytes was not due to an increase in neuronal fate. As Gfap can recognize both NSPCs and mature astrocytes, we employed Nestin and S100β to distinguish whether the decrease in oligodendrocytes we observed upon Nampt knockdown was due to a cell fate choice in these directions. While there was no detectable change in the generation of S100β+ mature astrocytes in Nampt Ad‐Cre cultures, there was a fourfold increase in the percentage of Nestin+ cells (6% in Nampt Ad‐LacZ cells; 23% in Nampt Ad‐Cre cells), suggesting quiescence rather than precocious astrocytic differentiation (Fig 5L). Importantly, all of these effects were completely rescued by treatment with NMN. However, we also observed a mild increase in TUNEL+ cell death under these conditions (33% increase relative to Nampt Ad‐LacZ cells), implying the possibility of a minor contribution of cell death to this phenotype. Together, these data suggest that genetic knockdown of Nampt prevents the successful differentiation of oligodendrocytes from NSPCs, potentially due to quiescence as indicated by the retention of Nestin staining.
Genetic knockdown of Nampt impairs OPC formation in vitro
Since differentiation using a non‐specific lineage differentiation protocol (by the removal of growth factors) revealed a specific requirement for Nampt in the successful generation of O4+ immature oligodendrocytes, we next asked which stage of NSPC differentiation into oligodendrocytes depends on Nampt by employing a differentiation protocol that promotes the oligodendrocyte lineage (Fig 6A, Supplementary Fig S5A). As previously observed using a non‐specific lineage differentiation protocol, the proportion of O4+ intermediate oligodendrocytes was dramatically decreased in Nampt Ad‐Cre cultures at 6–7 days post‐differentiation (Fig 6B). Furthermore, we observed that ablation of Nampt resulted in a decreased pool of OPCs (Pdgfrα+), but an increased pool of Nestin+ NSPCs. The proportion of Gfap+ astrocytes/NSPCs also mildly increased. To investigate whether the depletion of the OPC population was preexisting to or induced upon differentiation, we assessed the OPC population present during proliferation (Supplementary Fig S5B) and after 2 days of differentiation (Fig 6C), a time point that enriches for OPCs. Indeed, both of these time points also showed the loss of OPCs (Pdgfrα+, Olig2+). These results support our notion that Nampt is necessary for NSPCs to differentiate into OPCs.
How does Nampt promote oligodendrogenesis? Among NAD+‐consuming enzymes, the sirtuins Sirt1 and Sirt2 have been suggested to be involved in oligodendrogenesis (Li et al, 2007; Ji et al, 2011; Rafalski et al, 2013). Furthermore, we observed that Sirt2 was upregulated during oligodendrocyte differentiation in vitro and expressed in the SGZ in Nampt+ cells and NestinGFP+ NSPCs (Supplementary Fig S5C–F). To examine whether Sirt1 and Sirt2 play a role in Nampt‐mediated NSPC differentiation into oligodendrocytes, we acutely treated NSPCs with the selective inhibitor of Sirt2, AGK2, or the Sirt1 inhibitor, EX527. Surprisingly, whereas both inhibitors acutely suppressed oligodendrocyte formation (O4+, Fig 6D), neither chronic ablation of Sirt2 in the NSPCs isolated from Sirt2−/− mice nor Cre adenovirus‐mediated knockdown of Sirt1 in Sirt1flox/flox‐derived neurospheres affected oligodendrogenesis (Pdgfrα+, Olig2+, O4+), except that Sirt1 deficiency affected the production of O4+ intermediate oligodendrocytes (Fig 6E). Since Sirt1 and Sirt2 have multiple redundant targets (Luo et al, 2001; Jin et al, 2008; Rothgiesser et al, 2010), we speculated that they may have compensatory roles in the regulation of oligodendrogenesis. To test this possibility, we generated Sirt1/Sirt2‐double‐knockout (Sirt1/2 DKO) neurospheres. Consistent with the inhibitor studies (Fig 6D), dissociated Sirt1/2 DKO neurospheres were unable to form oligodendrocyte lineage cells upon differentiation (Fig 6E). To assess the role of Sirt1 and Sirt2 downstream of Nampt activity, we examined the expression of genes associated with OPC formation in Nampt Ad‐Cre neurospheres, Sirt1/2 DKO neurospheres, and their respective controls. Supporting our model, dissociated Nampt Ad‐Cre and Sirt1/2 DKO neurospheres showed similar decreases in the mRNA expression of Pdgfrα, Sox10, Nkx2.2 after 2 days of differentiation (Fig 6F and G). Moreover, dissociated Nampt Ad‐Cre and Sirt1/2 DKO neurospheres, respectively, exhibited similar increases in the expression of p21 (cdkn1a). Olig1 expression showed no change or slight reduction by these genetic ablations, potentially due to its lesser expression in NSPCs relative to Olig2 (Ligon et al, 2007) and predominant roles in oligodendrocyte maturation and remyelination rather than specification (Lu et al, 2002; Arnett et al, 2004). On the other hand, as previously observed (Saharan et al, 2013), neither Cre‐mediated knockdown of Sirt1 in neurospheres nor neurospheres cultured from whole‐body Sirt1−/− or Sirt2−/− mice exhibited defects in proliferation (Supplementary Fig S5G–J). Taken together, we conclude that Sirt1 and Sirt2 can redundantly mediate NSPC differentiation into OPCs.
Adult NSPC‐specific deletion of Nampt impairs NSPC differentiation in response to insult in vivo
While we observed a clear effect of Nampt ablation on NSPC differentiation into OPCs in vitro, we failed to detect a defect in oligodendrogenesis in the SGZ of iNSPC‐Nampt‐KO mice in vivo (Fig 3I), a finding consistent with previous reports that adult NSPCs rarely produce oligodendrocytes under basal conditions (Steiner et al, 2004). Since the SVZ is known to be more oligodendrogenic than the SGZ (Hack et al, 2004, 2005; Jackson et al, 2006; Menn et al, 2006), we assessed the percentage of NestinGFP+ cells that expressed Olig2 in the SVZs of iNSPC‐GFP and iNSPC‐Nampt‐KO mice (Fig 7A). Indeed, iNSPC‐Nampt‐KO mice showed a lower percentage of oligodendrocytes generated from adult NSPCs.
Given that aging impairs the ability of NSPCs to respond to environmental stimuli (Decker et al, 2002; Sim et al, 2002; Doucette et al, 2010), we hypothesized that Nampt‐mediated NAD+ biosynthesis is important to enable NSPCs to respond to such environmental perturbations. To test this idea, we employed the cuprizone model of demyelination and remyelination. Specifically, we fed 6‐ to 9‐week‐old iNSPC‐Nampt‐KO and littermate control mice (iNSPC‐GFP) a diet containing 0.2% cuprizone for 4–5 weeks, inducing deletion of Nampt in the adult Nestin+ population the week before starting the cuprizone diet (Fig 7B) (Skripuletz et al, 2011). To ensure that we were analyzing progeny of adult Nestin+ cells, all mice in our cohort expressed Cre recombinase under the inducible Nestin promoter (Nestin‐CreERT2) (Lagace et al, 2007) and the aforementioned Cre recombinase‐responsive GFP reporter transgene. Thus, we focused our analysis on lineage tracer marked (NestinGFP+) cells.
Cuprizone feeding did not alter the total number of NestinGFP+ cells present in the iNSPC‐GFP DG (Supplementary Fig S6A–C), suggesting that NSPC proliferation was unaltered. On the other hand, cuprizone‐fed mice exhibited an increased percentage of NestinGFP+ cells that colocalized with the NSPC markers Nestin+ (from 13 to 35%) and Gfap+ (from 19 to 41%), suggesting that cuprizone treatment prevented SGZ NSPCs from terminally differentiating and instead resulted in their retention of NSPC characteristics, which could occur through increased self‐renewal decisions and/or quiescence (Supplementary Fig S6D and E). To assess whether the NestinGFP‐marked NSPCs had differentiated into oligodendrocyte lineage cells in response to cuprizone, we next assessed colocalization between NestinGFP and oligodendrocyte‐specific markers, Sox10 and APC. However, the SGZ did not substantially produce oligodendrocytes even in response to demyelination (Supplementary Fig S6F and G).
While cuprizone treatment affects the hippocampus, its principle target of myelin injury is the corpus callosum (CC) (Gudi et al, 2009; Norkute et al, 2009; Doucette et al, 2010; Skripuletz et al, 2011). Under basal conditions and particularly during demyelination, NSPCs can proliferate to generate oligodendrocytes which integrate into the corpus callosum (Nait‐Oumesmar et al, 1999; Picard‐Riera et al, 2002; Hack et al, 2005; Menn et al, 2006; Colak et al, 2008; Jablonska et al, 2010; Soundarapandian et al, 2011). Thus, we assessed the fate decisions of migratory cells derived from the adult Nestin+ population in the subcallosal zone of the corpus callosum (Supplementary Fig S6A). In the iNSPC‐GFP CC, virtually no NestinGFP+ or Nestin+ cells were seen in regular chow‐fed mice (Fig 7C and D, Supplementary Fig S6A). There were no differences in the number of NestinGFP+ cells in the CC between control and iNSPC‐Nampt‐KO mice, suggesting that loss of Nampt neither affected insult‐induced NSPC proliferation or migration. In the iNSPC‐GFP CC, cuprizone feeding significantly increased the percentage of NestinGFP+Nestin+ cells (from 3 to 41%), but decreased the NestinGFP+Gfap+ (from 60 to 24%) double‐positive cells, suggesting increased self‐renewal fate decisions at the expense of astrocytic fate decisions (Fig 7E–G). In control mice, cuprizone feeding also increased the number of NestinGFP+ Sox10+ (from 2 to 16%) and NestinGFP+ Apc+ (from 0 to 4%) double‐positive cells, suggesting increased oligodendrocyte lineage fate decisions (Fig 7H and I). In contrast, the NestinGFP+ cells in the iNSPC‐Nampt‐KO CC showed significantly less colocalization with Nestin, Sox10, and Apc and more colocalization with Gfap (Fig 7F–I). Interestingly, Nampt was only expressed in the CC upon insult (Fig 7J, Supplementary Fig S6H). Moreover, Nampt colocalized with markers of NSPCs (Sox2, Supplementary Fig S6H) and oligodendrocytes (Olig2, Fig 7J). Together, these results suggest that Nampt is specifically expressed in SGZ/SVZ‐derived remyelinating NSPCs and plays an important role in oligodendrogenesis in response to insult.
Our study establishes a novel role for Nampt‐mediated NAD+ biosynthesis in NSPC proliferation, self‐renewal, and differentiation into oligodendrocytes. Importantly, we demonstrated that: (i) hippocampal NAD+ levels and Nampt expression decline with age; (ii) in vivo ablation of Nampt in the adult Nestin+ population impaired NSPC proliferation and self‐renewal; (iii) acute ablation of Nampt in hippocampal neurospheres significantly reduced NAD+ levels in NSPCs and stalled them in G1 of the cell cycle; (iv) chronic ablation of Nampt in hippocampal neurospheres abrogated oligodendrogenesis, and in vivo ablation of Nampt in the adult Nestin+ population reduced NSPC‐mediated oligodendrogenesis upon insult; (v) Sirt1 and Sirt2 are required for NSPC‐mediated oligodendrogenesis, yet in a redundant manner. These results reveal that Nampt deficiency in adult NSPCs recapitulates their functional defects observed during the aging process.
The finding that both hippocampal NAD+ levels and Nampt protein expression decline with age is consistent with the recent studies showing a significant age‐related decline in NAD+ and Nampt levels in multiple peripheral organs and tissues (Yoshino et al, 2011). Based on our findings in the present study, it is conceivable that loss of Nampt activity contributes to the age‐related decline in NSPC proliferation and in the NSPC pool (Lugert et al, 2010; Encinas et al, 2011; Artegiani & Calegari, 2012). This idea raises the possibility that long‐term NMN administration could combat age‐related declines in NSPC functionality. Indeed, we showed that the NSPC pool decreased with age and that long‐term NMN administration was able to maintain the NSPC pool, providing further support for the ‘depletion’ hypothesis of NSPC aging (Encinas et al, 2011; Artegiani & Calegari, 2012). However, the dosage of NMN that maintained the NSPC pool was not sufficient to drive the proliferation of aged NSPCs and only tended to increase neurogenesis itself. These results suggest that NAD+‐related effects on aged NSPCs are independent of the pathways altered by environmental enrichment (Kempermann et al, 1998, 2002) and exercise (Kronenberg et al, 2006; Wu et al, 2008), as these manipulations primarily affect NSPC proliferation and newborn neuron survival rather than the NSPC pool in aged animals. As we showed Nampt to be critically important for NSPC proliferation in neurospheres, it is likely that a higher dosage of NMN is necessary to promote NSPC proliferation. Yet, our results do not exclude the possibility that another factor present in the NSPC niche is more important for the age‐related decline in NSPC proliferation and/or counteracts Nampt/NMN‐driven NSPC proliferation. Importantly for therapeutic purposes, this experiment and our finding that intraperitoneal injection of NMN substantially increases hippocampal NAD+ levels within 15 min (Supplementary Fig S2G) strongly suggest that NMN can cross the blood–brain barrier.
It is remarkable that loss of Nampt in adult NSPCs produces functional impairment as hippocampal NSPCs could theoretically obtain NAD+ from release by surrounding cells. Studies to date have suggested that glia may deliver NAD+ to neurons (Verderio et al, 2001). Moreover, there are multiple precursors for and pathways of NAD+ biosynthesis, all of which are expressed in the hippocampus (Guillemin et al, 2007). Our results clearly demonstrate that cell‐autonomous Nampt activity is the main source of NAD+ for NSPCs. This cell‐autonomous Nampt activity is critical to regulate G1/S transition in NSPCs. Given that loss of Nampt activity in many other cell types reduced cell viability (Hasmann & Schemainda, 2003; Rongvaux et al, 2008), it is surprising that loss of Nampt activity specifically caused a G1/S stall without causing detectable cell death in NSPCs. As E2F1‐deficient mice have significantly reduced hippocampal NSPC death (Cooper‐Kuhn et al, 2002), the decrease in E2F1 expression that we observed upon inhibition of Nampt may explain this phenomenon (Fig 4E). In line with a G1/S stall, we also observed that loss of Nampt activity specifically downregulated cyclins E and A expression. E‐type cyclins regulate G1 progression, and their activity is necessary and limiting for the re‐entry of G0 cells into the cell cycle as well as for passing from G1 into S phase. Because we also observed downregulation of E2F1 expression, which transcriptionally regulates cyclin E, it is very likely that the downregulation of E2F1 contributes to the downregulation of cyclin E. Interestingly, we also see upregulation of p21 upon the loss of Nampt. p21 represses E2F activity and E2F activation is required to eliminate a p21‐mediated block in cell cycle entry in late G1 (Polager & Ginsberg, 2009; Wong et al, 2011). Thus, the upregulation of p21 that we see upon the loss of Nampt may also contribute to the downregulation of E2F/cyclin E activity.
Another important role of Nampt in NSPCs is to regulate their differentiation into oligodendrocytes. Why does loss of Nampt‐mediated NAD+ biosynthesis particularly affect oligodendrocytic lineage fate decisions in NSPCs? One possible explanation is that oligodendrocyte lineage cells are the most metabolically demanding (Sanchez‐Abarca et al, 2001) and stress‐sensitive central nervous system cell type (Husain & Juurlink, 1995). However, our results suggest that a more primary downstream effect of Nampt ablation on NSPC oligodendrocytic lineage fate decisions is the reduction in Sirt1 and Sirt2 activity (Fig 8). Previous work has generated contradictory conclusions in the function of Sirt1 in NSPC differentiation (Hisahara et al, 2008; Prozorovski et al, 2008; Wang et al, 2011b; Zhang et al, 2011; Rafalski et al, 2013; Saharan et al, 2013) and in the function of Sirt2 in oligodendrocyte differentiation (Li et al, 2007; Ji et al, 2011). Yet, these contradictory findings could be explained by compensatory activity between Sirt1 and Sirt2. Since Sirt1 is highly expressed in NSPCs (Hisahara et al, 2008; Prozorovski et al, 2008; Saharan et al, 2013) and Sirt2 is highly expressed in oligodendrocytes (Li et al, 2007; Tyler et al, 2011), it is possible that the activities of these two proteins trade off during oligodendrocyte maturation.
The factor downstream of Sirt1/2, which promotes oligodendrocytic fate decisions, remains to be identified. Previous work has linked Sirt1 activity to Hes1 (Hisahara et al, 2008), Hes5 (Hisahara et al, 2008), and Mash1 expression (Prozorovski et al, 2008; Zhang et al, 2011). Given the importance of Hes5 in inhibiting oligodendrocyte differentiation (Liu et al, 2006), it will be of great importance to investigate the roles of these factors downstream of Nampt. Our results reveal that ablation of Nampt specifically reduced the proportion of NSPC‐generated Pdgfrα+ OPCs as well as the transcription of Pdgfrα, Sox10, and Nkx2.2, but upregulated the expression of p21. One main regulator of these factors is the basic helix‐loop‐helix transcription factor Olig2 (Takebayashi et al, 2002; Zhou & Anderson, 2002; Ligon et al, 2007; Wegner, 2008). Olig2 is expressed in almost all (95–97%) of neurosphere cells and is necessary and sufficient to specify OPCs from the SVZ (Hack et al, 2004, 2005; Menn et al, 2006). Intriguingly, the number of Olig2+ cells in the SVZ decreases with age (Bouab et al, 2011). Recent work has shown that Olig2 competes with p53 for binding to an important coregulator that regulates p53 acetylation (Mehta et al, 2011; Sun et al, 2011). As Sirt1 and Sirt2 can deacetylate p53 (Polager & Ginsberg, 2009; van Leeuwen et al, 2013), it will be interesting to investigate the NAD+ dependence of Olig2 activity.
Identifying Nampt as a critical regulator of NSPC proliferation, self‐renewal, and differentiation highlights the intimate link between stem cell metabolism and cell fate (Folmes et al, 2012; Zhang et al, 2012). Like loss of Nampt, inhibition of mitochondrial fatty acid oxidation impairs stem cell self‐renewal decisions (Ito et al, 2012) and abrogation of de novo lipogenesis impairs NSPC proliferation (Knobloch et al, 2013). However, manipulations of other metabolic pathways can present very differently from loss of Nampt. For example, ablation of endogenous cholesterol production in NSPCs results in massive apoptosis of newborn neurons without affecting cell death or proliferation in the NSPCs themselves (Saito et al, 2009). And, mitochondrial inhibition alters NSPC neuronal lineage fate decisions without affecting the astrocytic or oligodendrocytic lineages (Voloboueva et al, 2010). Thus, our work supports the notion that cellular metabolism regulates NSPC functionality, but that alteration of each metabolic pathway generates specific effects.
Aging is an opposing factor for any type of regeneration (Jadasz et al, 2012). Consistent with the notion that altered metabolism plays a causal role in the age‐related decline in NSPC function, NSPCs isolated from old mice had decreased regenerative capacity, fewer functional mitochondria, and less oxygen consumption relative to NSPCs isolated from young mice (Stoll et al, 2011). Rates of remyelination decline with age due to slower recruitment of OPCs into areas of demyelination and slower differentiation of OPCs into remyelinating oligodendrocytes (Decker et al, 2002; Sim et al, 2002; Doucette et al, 2010). Similarly, the lack of remyelination seen in multiple sclerosis, an autoimmune disease that leads to oligodendrocyte loss and demyelination (Jadasz et al, 2012), occurs with a failure of OPC differentiation (Franklin & Ffrench‐Constant, 2008). Our results clearly showed that in neurospheres, treatment with NMN rescued defects in oligodendrogenesis caused by a reduction in NAD+ levels. Furthermore, systemic NMN administration was able to substantially augment hippocampal NAD+ levels and increase the NSPC pool. Thus, NMN administration could be an efficient intervention to enhance the NSPC pool and promote remyelination by activating endogenous NSPCs during the aging process and/or in neurodegenerative diseases that cause demyelination. In conclusion, our present study provides impetus to further delineate the therapeutic potential and signaling pathways downstream of Nampt‐mediated NSPC self‐renewal, proliferation, and differentiation into oligodendrocytes.
Materials and Methods
Mice were maintained on a regular chow ad libitum on a 12 h light/dark cycle (lights on from 6 am to 6 pm). Namptflox/flox mice (Rongvaux et al, 2008), in which exons 5 and 6 of the Nampt gene are flanked by loxP sites, were crossed to Nestin‐CreERT2 mice (Lagace et al, 2007) to generate Nampt flox/+; Cre double heterozygous mice. Double heterozygous mice were bred to Namptflox/flox mice to obtain Namptflox/flox; Cre mutant mice (iNSPC‐Nampt‐KO mice) in the expected Mendelian ratio. To trace the progeny of adult NSPCs and to confirm the specificity and magnitude of the recombination induced by tamoxifen injection, iNSPC‐Nampt‐KO and Nestin‐CreERT2 mice were crossed to a reporter mouse strain that expresses a loxP‐flanked STOP cassette that prevents transcription of the downstream enhanced green fluorescent protein [ZsGreen1; Jackson laboratories #7906 (Madisen et al, 2010)]. Recombination PCR on hippocampal extracts of tamoxifen‐ or vehicle‐treated mice showed successful deletion upon treatment with tamoxifen (Supplementary Fig S2C). All animal procedures were approved by the Washington University Animal Studies Committee and were in accordance with NIH guidelines.
Induction of Nampt deletion
Tamoxifen injections were performed as described previously (Lagace et al, 2007). Briefly, iNSPC‐Nampt‐KO mice (5–7 weeks old) were administered tamoxifen (TAM, Sigma T5648) at 180 mg/kg/d for 5 days (d, intraperitoneally; dissolved in 10% EtOH/90% sunflower oil), a protocol that produces maximal recombination with minimal lethality (5%) (Lagace et al, 2007).
5′‐Bromodeoxyuridine (BrdU, Sigma, B9285) was diluted in sterile saline and administered by intraperitoneal injections (100 mg/kg body weight). For analysis of the cumulative effects of loss of Nampt, mice were given BrdU twice a day for 2 days and sacrificed the following day or 28 days later. For analysis of the effect of loss of Nampt on adult NSC differentiation and postnatal oligodendrocyte differentiation, mice were given BrdU twice a day for 1 day and sacrificed 2 days later.
Demyelination was induced by feeding 6‐ to 8‐week‐old mice a diet containing 0.2% cuprizone (bis‐cyclohexanone oxaldihydrazone; Sigma C9012) mixed into a ground standard rodent chow for 4–5 weeks (Harlan Laboratories, TD.01453). To allow recovery from cuprizone treatment, food was replaced with standard chow for an additional 1 week. This protocol has been shown to successfully demyelinate and remyelinate the hippocampus (Skripuletz et al, 2011).
The following primary and secondary antibodies were used:
Primary antibodies and their uses or cell type specificities (von Bohlen und Halbach, 2011):
Actin: normalization, WB 1:4,000 CPO1, Sigma; Gapdh: normalization, WB 1:4,000 6C5 Millipore CB1001; Nampt: IHC 1:1,000; WB 1:3,000 Alexis Biochemicals ALX‐804‐717‐C100; Pdgfrα: oligodendrocyte precursor cells, IF 1:500 APA5 BD Biosciences; Olig2: all oligodendrocyte lineage cells, IHC 1:500, IF 1:1,000; Millipore; O4: immature oligodendrocytes, IF 1:1,000 Millipore, MAB345; APC: oligodendrocytes, IHC 1:1,000 Millipore CC‐1 OP80; MBP: mature oligodendrocytes, IHC 1:1,000 Millipore MAB386; Ki67: proliferating cells, IHC, IF 1:500; WB 1:3,000 Abcam ab66155; Pcna: proliferating cells, WB 1:2,000; PC10 Cell signaling #2586; 5‐bromo‐2′‐deoxyuridine (BrdU): a thymine analog that incorporates into the DNA of cells in S phase, IHC 1:500; OBT0030 Accurate; activated caspase‐3: apoptosis, IHC, IF 1:500; Cell Signaling #9661; LC3B: autophagy, WB 1:1,000; Novus NB600‐1384; TUNEL: cell death, Roche In Situ Cell Death Detection Kit 11 684 795 910; Dcx: newly born neurons, IHC 1:1,000; Cell Signaling #4604; NeuN: mature neurons, IHC, 1:500, Millipore, MAB377; Nestin: NSPCs, IHC, IF 1:1,000, Millipore MAB353; Sox2: NSPCs, IHC, IF 1:500; WB 1:2,000; Millipore AB5603; Gfap: NSPCs and astrocytes, IHC, IF 1:1,000; Millipore MAB360.
Secondary antibodies: Jackson ImmunoResearch anti‐rat, anti‐rabbit, anti‐mouse Cy3 (1:400), Alexa Fluor488 (1:200), and Alexa Fluor647 (1:200). Anti‐rabbit, anti‐mouse horseradish peroxidase (Invitrogen).
FK866 (Hasmann & Schemainda, 2003) (Sigma F8557), EX527 (Peck et al, 2010) (Cayman Chemical 10009798), and AGK2 (Outeiro et al, 2007) (Sigma A8231) were dissolved in DMSO and used to inhibit Nampt, Sirt1, and Sirt2, respectively.
Neurosphere cultures and culture media were prepared as described previously with minor modifications (Dasgupta & Gutmann, 2005; Lu & Ramanan, 2012). Briefly, postnatal hippocampi were dissected in Hibernate A (Invitrogen, A12475‐01) and trypsinized at 37°C for 7 min. Cells were mechanically dissociated by pipetting and pelleted by centrifugation (213 g, 7 min). Dissociation medium (0.1% sodium bicarbonate, 15 mM HEPES, 0.5% glucose in HBSS) was used to wash the cells before they were resuspended in growth medium. Growth medium consisted of DMEM:F12 (1:1, Invitrogen 11966‐025 and 21700‐075, respectively), B27 (Invitrogen, 17504‐044), N2 (Invitrogen, 17502‐048), Pen/Strep (Invitrogen), epidermal growth factor (EGF, 20 ng/ml, Sigma, E4127), fibroblast growth factor (FGF, 10 ng/ml, R&D Systems, 233‐fb), and heparin (Sigma). Cultures were maintained at 37°C with 5% CO2 and passaged twice before use in experiments. Three to nine independent samples, each in 1–3 replicates, from at least two different litters were used in all experiments. Neurospheres were cultured in the physiological glucose level of 5 mM (Dienel & Cruz, 2006), which has been previously shown to have no negative consequences on NSPC proliferation, differentiation, or death (Fu et al, 2006; Gao & Gao, 2007).
Neurospheres derived from Namptflox/flox mice were infected with Ad5 Cre recombinase‐ or β‐galactosidase‐expressing (LacZ, control) adenoviruses at an MOI of 100. All assessments were performed at least 6 days post‐infection.
Neurosphere proliferation analysis
Neurospheres derived from Namptflox/flox mice were dissociated by trypsin digestion and seeded at similar cell densities in 24‐well plates with fresh growth medium. Every 24 h, neurospheres from triplicate wells were collected, dissociated, and counted on a hemocytometer using 0.2% trypan blue exclusion to distinguish viable cells. For analysis of neurosphere diameter, the largest neurosphere in each well was imaged (20× objective) and the diameter was calculated using ImageJ. For secondary neurosphere analysis, the total number of neurospheres in each well was counted at 7 days post‐plating.
Three to five days after their first passage, neurospheres were trypsinized, washed with dissociation medium, and plated at 150,000 cells per well in 24‐well plates in differentiation medium [growth medium without FGF and EGF and with BDNF (5 ng/ml, Peprotech, 450‐02) on glass coverslips coated with poly‐D‐lysine (50 μg/ml; Sigma) and laminin (20 μg/ml; BD Biosciences)]. Six‐well plates were coated with poly‐D‐lysine (20 μg/ml) and laminin (10 μg/ml). To enrich for oligodendrocytes, PDGFαα (10 ng/ml, Peprotech 100‐13A) was added to neurospheres at passage 2 and PDGFαα (2.5 ng/ml) and 3,3′,5‐triiodo‐L‐thyronine (T3, 40 ng/ml, Sigma T4397) were added to differentiation medium. The percentage of oligodendrocyte precursor cells (OPCs) generated was analyzed after 2 days of differentiation, and the percentage of differentiated oligodendrocytes was analyzed after 6–7 days of differentiation.
All tissue sections and cells were incubated in blocking/permeabilization solution containing 10% normal goat serum, 1% BSA, and 0.3% Triton X‐100 in PBS for 45–60 min prior to 24 or 48 h of incubation with primary antibodies in 5% normal goat serum and 0.1% Triton X‐100 in PBS at 4°C at the concentrations listed below. Alexa627‐, Alexa488‐, or Cy3‐conjugated secondary antibodies diluted in 2% normal goat serum, 1% BSA, and 0.1% Triton X‐100 in PBS were added for 2 h at room temperature. Nuclei were stained with 4,6‐diamidino‐2‐phenylindole (Sigma) for 10 min at room temperature.
Cells were harvested by fixation with 4% paraformaldehyde in PBS (15 min). Mice were anesthetized by i.p. injection of ketamine and xylazine and perfused transcardially through left ventricle with cold 0.1 M phosphate buffer at pH 7.4 followed by a phosphate‐buffered solution of 4% paraformaldehyde (PFA). Brains were post‐fixed with 4% PFA overnight and placed into 15% sucrose followed by 30% sucrose, frozen, and stored at −80°C until use. Coronal sections (30 μm) were made by cryostat in a 1 in 8 series and stored at −30°C in cryoprotectant until use. To remove any endogenous peroxidase activity, all sections were incubated with 3% H2O2 for 10 min. Tissue sections used to assess BrdU incorporation were treated before the immunostaining procedure with 50% formamide in 2× saline/sodium citrate (SSC) at 65°C for 2 h, 2N HCl for 30 min at 37°C, 0.1 M borate pH 8.5, and then washed twice with PBS before proceeding with the staining protocol. Tissue sections not used to assess BrdU incorporation were either incubated in 50% formamide in 2× saline/sodium citrate (SSC) at 65°C for 2 h or 10 mM citrate buffer at 65°C for 1 h before proceeding with the staining protocol. Detection of Dcx, Nestin, Nampt, and APC was performed using the TSA‐Plus fluorescein kit (PerkinElmer).
For tissue sections, high‐magnification (20×, 0.8DICII or 40× oil 1. 3DICII) microscopic imaging was performed using a Zeiss Axioimager.Z1. Images were taken in z‐stacks of 1 μm steps through the range of tissue section immunoreactivity. For the dorsolateral corner of the SVZ, images were taken from bregma 1.10 to −0.10 mm. For the corpus callosum, images were taken from bregma −1.06 to −2.54 mm. For the dentate gyrus, images were taken from bregma −1.34 to −3.64 mm. Quantification was performed blinded to genotype on 3–8 tissue sections per animal. Cell densities were estimated by the number of immunoreactive cells divided by the area of the structure, measured with ImageJ. Verification of colocalization was achieved by importing stacks of Z images into ImageJ and performing 3D rendering. For cells, 10 or 20× microscopic imaging was performed using a Zeiss Axioimager.Z1. Quantification was performed blinded to genotype on 2–3 fields of view per sample and treatment, from 3 to 9 independent samples.
NAD+ levels were determined using an HPLC system (Shimadzu) with a Supelco LC‐18‐T column (15 × 4.6 cm; Sigma), as described previously (Yoshino et al, 2011).
Microarrays and bioinformatic analyses
For individual genes, raw microarray data were subjected to Z score transformation, and Z ratios were calculated as described previously (Cheadle et al, 2003). Subsequent analysis, namely parametric analysis of gene set enrichment (PAGE), was performed as previously described (Yoshino et al, 2011). The microarray data used in this study have been deposited into the NCBI GEO database (GEO accession number GSE49784).
Protein extracts (15–50 μg) from mouse hippocampi or neurospheres were prepared as previously described (Yoshino et al, 2011).
Quantitative real‐time RT–PCR
Total RNA was extracted from the hippocampus using the RNeasy kit (Qiagen) and reverse‐transcribed into cDNA with the High Capacity cDNA Reverse Transcription kit (Applied Biosystems). Quantitative real‐time RT–PCR was conducted with the TaqMan Fast Universal PCR Master mix and appropriate TaqMan primers for each gene with the GeneAmp 7500 fast sequence detection system (Applied Biosystems). Relative expression levels were calculated for each gene by normalizing to Gapdh levels and then to a control.
Differences between the two groups were assessed using the Student's unpaired t‐test. Comparisons among several groups were performed using one‐way ANOVA with the Tukey–Kramer post hoc test except for Fig 3J and Supplementary Fig S3D and E, in which the Games–Howell post hoc test and the Fisher LSD post hoc test were used, respectively. P‐values < 0.05 were considered statistically significant.
LRS and SI designed research, analyzed data, and wrote the paper. LRS performed research.
Conflict of interest
S.I. served as a scientific advisory board member for Sirtris, a GSK company, and had a sponsored research agreement with Oriental Yeast Co., Tokyo, Japan. S.I. is also a co‐founder of Metro Midwest Biotech.
Supplementary Figure S1
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We thank Jun Yoshino, Da Yong Lee, Paul P. Y. Lu, and members of the Imai laboratory for technical help and advice. We thank David Gutmann, Narendrakumar Ramanan, and Kelly Monk for making suggestions on and critically reading this manuscript. We thank the Oriental Yeast Co. (Tokyo, Japan) for providing the NMN used in the long‐term NMN administration study. This work was supported in part by the National Institute on Aging (AG024150, AG037457) and the Ellison Medical Foundation to S.I, and Training Grant T32 GM007067 to L.R.S.
FundingNational Institute on Aging AG024150 AG037457
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